Advances in Food Mycology

Advances in Experimental Medicine and Biology Editorial Board: NATHAN BACK, State University of New York at Buffalo IRUN R. COHEN, The Weizmann Institute of Science DAVID KRITCHEVSKY, Wistar Institute ABEL LAJTHA, N.S. Kline Institute for Psychiatric Research RODOLFO PAOLETTI, University of Milan Recent Volumes in this Series Volume 563 UPDATES IN PATHOLOGY Edited by David C. Chhieng and Gene P. Siegal Volume 564 GLYCOBIOLOGY AND MEDICINE: PROCEEDINGS OF THE 7TH JENNER GLYCOBIOLOGY AND MEDICINE SYMPOSIUM Edited by John S. Axford Volume 565 SLIDING FILAMENT MECHANISM IN MUSCLE CONTRACTION: FIFTY YEARS OF RESEARCH Edited by Haruo Sugi Volume 566 OXYGEN TRANSPORT TO ISSUE XXVI Edited by Paul Okunieff, Jacqueline Williams, and Yuhchyau Chen Volume 567 THE GROWTH HORMONE-INSULIN-LIKE GROWTH FACTOR AXIS DURING DEVELOPMENT Edited by Isabel Varela-Nieto and Julie A. Chowen Volume 568 HOT TOPICS IN INFECTION AND IMMUNITY IN CHILDREN II Edited by Andrew J. Pollard and Adam Finn Volume 569 EARLY NUTRITION AND ITS LATER CONSEQUENCES: NEW OPPORTUNITIES Edited by Berthold Koletzko, Peter Dodds, Hans Akerbloom, and Margaret Ashwell Volume 570 GENOME INSTABILITY IN CANCER DEVELOPMENT Edited by Erich A. Nigg Volume 571 ADVANCES IN MYCOLOGY Edited by J.I. Pitts, A.D. Hocking, and U. Thrane A Continuation Order Plan is available for this series. A continuation order will bring delivery of each new volume immediately upon publication. Volumes are billed only upon actual shipment. For further information please contact the publisher.

Advances in Food Mycology Edited by

A.D. Hocking Food Science Australia North Ryde, Australia

J.I. Pitt Food Science Australia North Ryde, Australia

R.A. Samson Centraalbureau voor Schimmelcultures Utrecht, Netherlands

and

U. Thrane Technical University of Denmark Lyngby, Denmark

A.D. Hocking Food Science Australia PO Box 52, North Ryde NSW 1670 Australia [email protected]

J.I. Pitt Food Science Australia PO Box 52, North Ryde NSW 1670 Australia [email protected]

R.A. Samson Centraalbureau voor Schimmelcultures PO Box 85167 3508 AD Utrecht Netherlands [email protected]

U. Thrane Biocentrum-DTU Technical University of Denmark Building 221 DK-2800 Kgs. Lyngby Denmark [email protected]

Library of Congress Control Number: 2005930810 ISBN-10: 0-387-28385-4 ISBN-13: 978-0387-28385-2

e-ISBN: 0-387-28391-9

Printed on acid-free paper. © 2006 Springer Science+Business Media, Inc. All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, Inc., 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed in the United States of America. 9 8 7 6 5 4 3 2 1 springeronline.com

(SPI/EB)

FOREWORD

This book represents the Proceedings of the Fifth International Workshop on Food Mycology, which was held on the Danish island of Samsø from 15-19 October, 2003. This series of Workshops commenced in Boston, USA, in July 1984, from which the proceedings were published as Methods for Mycological Examination of Food (edited by A. D. King et al., published by Plenum Press, New York, 1986). The second Workshop was held in Baarn, the Netherlands, in August 1990, and the proceedings were published as Modern Methods in Food Mycology (edited by R. A. Samson et al., and published by Elsevier, Amsterdam, 1992). The Third Workshop was held in Copenhagen, Denmark, in 1994 and the Fourth near Uppsala, Sweden, in 1998. The proceedings of those two workshops were published as scientific papers in the International Journal of Food Microbiology. International Workshops on Food Mycology are held under the auspices of the International Commission on Food Mycology, a Commission under the Mycology Division of the International Union of Microbiological Societies. Details of this Commission are given in the final chapter of this book. This Fifth Workshop was organised by Ulf Thrane, Jens Frisvad, Per V. Nielsen and Birgitte Andersen from the Center for Microbial Biotechnology, Technical University of Denmark, Kgs. Lyngby,

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Foreword

Denmark. This Center, through numerous publications and both undergraduate teaching and graduate supervision has been highly influential in the world of food mycology for the past 20 years and more. Trine Bro and Lene Nordsmark from the Center also carried out the important tasks of providing secretarial help to the Organisers and solving the logistics of moving participants from Copenhagen to Samsø and back. Samsø provided an ideal setting for the Fifth Workshop, as the island is made up of rural agricultural communities, with old villages and rustic land and seascapes. The Fifth Workshop was attended by some 35 participants, drawn from among food mycology and related disciplines around the world. The workshop was highly successful, with papers devoted to media and methods development in food mycology, as is usual with these workshops. Particular emphasis was placed on the fungi which produce mycotoxins, especially their ecology, and through ecology, potential control measures. Sessions were also devoted to yeasts, and the inactivation of fungal spores by the use of heat and high pressure. Nearly 40 scientific papers were presented over three days of the workshop, and these papers are the major contributions in these Proceedings. The organisers especially wish to thank the sponsors of the Fifth Workshop: BCN Laboratories, Knoxville, Tennessee, USA; the Danish ECB5 Foundation, Copenhagen; Novozymes A/S, Bagsværd, Denmark; LMC Centre for Advanced Food Studies, Copenhagen; the Danish Research Agency STVF, Copenhagen, though Grant Number 26-03-0188; Eurofins Denmark A/S, Copenhagen and the Mycology Division of the International Union of Microbiological Societies, for their support which made this workshop possible. A.D. Hocking J.I. Pitt R.A. Samson U. Thrane

CONTENTS

Foreword . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xi Section 1. Understanding the fungi producing important mycotoxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Important mycotoxins and the fungi which produce them . . . . . . Jens C. Frisvad, Ulf Thrane, Robert A. Samson and John I. Pitt Recommendations concerning the chronic problem of misidentification of mycotoxigenic fungi associated with foods and feeds . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jens C. Frisvad, Kristian F. Nielsen and Robert A. Samson

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Section 2. Media and method development in food mycology . . . .

47

Comparison of hyphal length, ergosterol, mycelium dry weight, and colony diameter for quantifying growth of fungi from foods . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marta H. Taniwaki, John I. Pitt, Ailsa D. Hocking and Graham H. Fleet

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Contents

Evaluation of molecular methods for the analysis of yeasts in foods and beverages . . . . . . . . . . . . . . . . . . . . . . . . . . Ai Lin Beh, Graham H. Fleet, C. Prakitchaiwattana and Gillian M. Heard

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Standardization of methods for detecting heat resistant fungi . . 107 Jos Houbraken and Robert A. Samson Section 3. Physiology and ecology of mycotoxigenic fungi . . . . . . 113 Ecophysiology of fumonisin producers in Fusarium section Liseola . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Vicente Sanchis, Sonia Marín, Naresh Magan and Antonio J. Ramos Ecophysiology of Fusarium culmorum and mycotoxin production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123 Naresh Magan, Russell Hope and David Aldred Food-borne fungi in fruit and cereals and their production of mycotoxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 Birgitte Andersen and Ulf Thrane Black Aspergillus species in Australian vineyards: from soil to ochratoxin A in wine . . . . . . . . . . . . . . . . . . . . . . . . . 153 Su-lin L. Leong, Ailsa D. Hocking, John I. Pitt, Benozir A. Kazi, Robert W. Emmett and Eileen S. Scott Ochratoxin A producing fungi from Spanish vineyards . . . . . . . 173 Marta Bau, M. Rosa Bragulat, M. Lourdes Abarca, Santiago Minguez and F. Javier Cabañes Fungi producing ochratoxin in dried fruits . . . . . . . . . . . . . . . . Beatriz T. Iamanaka, Marta H. Taniwaki, E. Vicente and Hilary C. Menezes

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An update on ochratoxigenic fungi and ochratoxin A in coffee . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 Marta H. Taniwaki

Contents

Mycobiota, mycotoxigenic fungi, and citrinin production in black olives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dilek Heperkan, Burçak E. Meriç, Gülçin Sismanoglu, Gözde Dalkiliç and Funda K. Güler

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Byssochlamys: significance of heat resistance and mycotoxin production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jos Houbraken, Robert A. Samson and Jens C. Frisvad

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Effect of water activity and temperature on production of aflatoxin and cyclopiazonic acid by Aspergillus flavus in peanuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Graciela Vaamonde, Andrea Patriarca and Virginia E. Fernández Pinto

225

Section 4. Control of fungi and mycotoxins in foods . . . . . . . . . . 237 Inactivation of fruit spoilage yeasts and moulds using high pressure processing . . . . . . . . . . . . . . . . . . . . . . . . Ailsa D. Hocking, Mariam Begum and Cindy M. Stewart

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Activation of ascospores by novel food preservation techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jan Dijksterhuis and Robert A. Samson

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Mixtures of natural and synthetic antifungal agents . . . . . . . . . Aurelio López-Malo, Enrique Palou, Reyna León-Cruz and Stella M. Alzamora

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Probabilistic modelling of Aspergillus growth . . . . . . . . . . . . . . Enrique Palou and Aurelio López-Malo

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Antifungal activity of sourdough bread cultures . . . . . . . . . . . . 307 Lloyd B. Bullerman, Marketa Giesova, Yousef Hassan, Dwayne Deibert and Dojin Ryu Prevention of ochratoxin A in cereals in Europe . . . . . . . . . . . . 317 Monica Olsen, Nils Jonsson, Naresh Magan, John Banks, Corrado Fanelli, Aldo Rizzo, Auli Haikara, Alan Dobson, Jens Frisvad, Stephen Holmes, Juhani Olkku, Sven-Johan Persson and Thomas Börjesson

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Contents

Recommended methods for food mycology . . . . . . . . . . . . . . .

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Appendix 1 – Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Appendix 2 – International Commission on Food Mycology . .

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Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 361

CONTRIBUTORS

M. Lourdes Abarca, Departament de Sanitat i d’Anatomia Animals, Universitat Autónoma de Barcelona, 08193 Bellaterra, Barcelona, Spain David Aldred, Applied Mycology Group, Biotechnology Centre, Cranfield University, Silsoe, Bedford MK45 4DT, UK Stella M. Alzamora, Departament de Industrias, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Universitaria 1428, Buenos Aires, Argentina Birgitte Andersen, Center for Microbial Biotechnology, BioCentrumDTU, Technical University of Denmark, DK-2800 Kgs. Lyngby, Denmark Marta Bau, Departament de Sanitat i d’Anatomia Animals, Universitat Autónoma de Barcelona, 08193 Bellaterra, Barcelona, Spain John Banks, Central Science Laboratory, Sand Hutton, York YO41 1LZ, UK Mariam Begum, Food Science Australia, CSIRO, P.O. Box 52, North Ryde, NSW 1670, Australia Ai Lin Beh, Food Science and Technology, School of Chemical Engineering and Industrial Chemistry, University of New South Wales, Sydney, NSW 2052, Australia Thomas Börjesson, Svenska Lantmännen, Östra hamnen, SE-531 87 Lidköping, Sweden

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Contributors

M. Rosa Bragulat, Departament de Sanitat i d’Anatomia Animals, Universitat Autónoma de Barcelona, 08193 Bellaterra, Barcelona, Spain Lloyd B. Bullerman, Department of Food Science and Technology, University of Nebraska, Lincoln, NE 68583-0919, USA F. Javier Cabañes, Departament de Sanitat i d’Anatomia Animals, Universitat Autónoma de Barcelona, 08193 Bellaterra, Barcelona, Spain Gözde Dalkiliç, Department of Food Engineering, Istanbul Technical University, Istanbul, 34469 Maslak, Turkey Dwayne Deibert, Department of Food Science and Technology, University of Nebraska, Lincoln, NE 68583-0919, USA Jan Dijksterhuis, Department of Applied Research and Services, Centraalbureau voor Schimmelcultures, Fungal Biodiversity Centre, Uppsalalaan 8, 3584 CT, Utrecht, Netherlands Alan Dobson, Microbiology Department, University College Cork, Cork, Ireland Robert W. Emmett, Department of Primary Industries, PO Box 905, Mildura, Vic. 3502, Australia Corrado Fanelli, Laboratorio di Micologia, Univerisità “La Sapienza”, Largo Cristina di Svezia 24, I-00165 Roma, Italy Virginia E. Fernández Pinto, Laboratorio de Microbiología de Alimentos, Departamento de Química Orgánica, Area Bromatología, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Universitaria, Pabellón II, 3˚ Piso, 1428, Buenos Aires, Argentina Graham H. Fleet, Food Science and Technology, School of Chemical Engineering and Industrial Chemistry, University of New South Wales, Sydney, NSW 2052, Australia Jens C. Frisvad, BioCentrum-DTU, Building 221, Technical University of Denmark, 2800 Lyngby, Denmark Marketa Giesova, Department of Dairy and Fat Technology, Institute of Chemical Technology, Prague, Czech Republic Funda K. Güler Istanbul Technical University, Dept. of Food Engineering Istanbul, Turkey, 34469 Maslak Auli Haikara, VTT Biotechnology, PO Box 1500, FIN-02044 Espoo, Finland Yousef Hassan, Department of Food Science and Technology, University of Nebraska, Lincoln, NE 68583-0919, USA Gillian M. Heard, Food Science and Technology, School of Chemical Engineering and Industrial Chemistry, University of New South Wales, Sydney, NSW 2052, Australia

Contributors

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Dilek Heperkan, Istanbul Technical University, Dept. of Food Engineering Istanbul, 34469 Maslak, Turkey Ailsa D. Hocking, Food Science Australia, CSIRO, PO Box 52, North Ryde, NSW 1670, Australia Stephen Holmes, ADGEN Ltd, Nellies Gate, Auchincruive, Ayr KA6 5HW, UK Russell Hope, Applied Mycology Group, Biotechnology Centre, Cranfield University, Silsoe, Bedford MK45 4DT, UK Jos Houbraken, Centraalbureau voor Schimmelcultures, PO Box 85167, 3508 AD, Utrecht, The Netherlands Beatriz T. Iamanaka, Food Technology Institute, ITAL C.P 139 CEP13.073-001 Campinas-SP, Brazil Nils Jonsson, Swedish Institute of Agricultural and Environmental Engineering, PO Box 7033, SE-750 07 Uppsala, Sweden Benozir A. Kazi, Department of Primary Industries, PO Box 905, Mildura, Vic. 3502, Australia Reyna León-Cruz, Ingeniería Química y Alimentos, Universidad de las Américas, Puebla, Cholula 72820, Mexico Su-lin L. Leong, Food Science Australia, CSIRO, PO Box 52, North Ryde, NSW 1670, Australia Aurelio López-Malo, Ingeniería Química y Alimentos, Universidad de las Américas, Puebla, Cholula 72820, Mexico Naresh Magan, Applied Mycology Group, Biotechnology Centre, Cranfield University, Barton Road, Silsoe, Bedford MK45 4DT, UK Sonia Marín, Food Technology Department, Lleida University, 25198 Lleida, Spain Hilary C. Menezes, Food Engineering Faculty (FEA), Unicamp, Campinas-SP, Brazil Burçak E. Meriç, Istanbul Technical University, Dept. of Food Engineering Istanbul, 34469 Maslak, Turkey Santiago Minués, Institut Català de la Vinya i el Vi (INCAVI), Generalitat de Catalunya, Vilafranca del Penedés, Barcelona, Spain Kristian F. Nielsen, BioCentrum-DTU, Building 221, Technical University of Denmark, 2800 Lyngby, Denmark Juhani Olkku, Oy Panimolaboratorio-Bryggerilaboratorium AB, P.O. Box 16, FIN-02150 Espoo, Finland Monica Olsen, National Food Administration, PO Box 622, SE-751 26 Uppsala, Sweden Enrique Palou, Ingeniería Química y Alimentos, Universidad de las Américas, Puebla, Cholula 72820, Mexico

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Contributors

Andrea Patriarca, Laboratorio de Microbiología de Alimentos, Departamento de Química Orgánica, Area Bromatología, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Universitaria, Pabellón II, 3˚ Piso, 1428, Buenos Aires, Argentina Sven-Johan Persson, Akron maskiner, SE-531 04 Järpås, Sweden John I. Pitt, Food Science Australia, CSIRO, PO Box 52, North Ryde, NSW 1670, Australia C. Prakitchaiwattana, Food Science and Technology, School of Chemical Engineering and Industrial Chemistry, University of New South Wales, Sydney, NSW 2052, Australia Antonio J. Ramos, Food Technology Department, Lleida University, 25198 Lleida, Spain Aldo Rizzo, National Veterinary and Food Res. Inst., PO Box 45, FIN-00581, Helsinki, Finland Dojin Ryu, Department of Food Science and Technology, University of Nebraska, Lincoln, NE 68583-0919, USA Robert A. Samson, Department of Applied Research and Services, Centraalbureau voor Schimmelcultures, Fungal Biodiversity Centre, Uppsalalaan 8, 3584 CT, Utrecht, Netherlands Vicente Sanchis, Food Technology Department, Lleida University, 25198 Lleida, Spain Eileen S. Scott, School of Agriculture and Wine, University of Adelaide, PMB 1, Glen Osmond, SA 5064, Australia Gülçin S¸is¸manog˘lu, Department of Food Engineering, Istanbul Technical University, Istanbul, 34469 Maslak, Turkey Cindy Stewart, National Center for Food Safety and Technology, 6502 S. Archer Rd, Summit-Argo, IL 60501, USA Marta H. Taniwaki, Food Technology Institute (ITAL), C.P 139 CEP13.073-001 Campinas-SP, Brazil Ulf Thrane, Center for Microbial Biotechnology, BioCentrum-DTU, Technical University of Denmark, DK-2800 Kgs. Lyngby, Denmark Graciela Vaamonde, Laboratorio de Microbiología de Alimentos, Departamento de Química Orgánica, Area Bromatología, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Universitaria, Pabellón II, 3˚ Piso, 1428, Buenos Aires, Argentina E. Vicente Food Technology Institute, ITAL C.P 139 CEP13.073-001 Campinas-SP, Brazil

Section 1. Understanding the fungi producing important mycotoxins Important mycotoxins and the fungi which produce them Jens C. Frisvad, Ulf Thrane, Robert A. Samson and John I. Pitt Recommendations concerning the chronic problem of misidentification of mycotoxigenic fungi associated with foods and feeds Jens C. Frisvad, Kristian F. Nielsen and Robert A. Samson

IMPORTANT MYCOTOXINS AND THE FUNGI WHICH PRODUCE THEM Jens C. Frisvad, Ulf Thrane,* Robert A. Samson† and John I. Pitt‡

1.

INTRODUCTION

The assessment of the relationship between species and mycotoxins production has proven to be very difficult. The modern literature is cluttered with examples of species purported to make particular mycotoxins, but where the association is incorrect. In some cases, mycotoxins have even been named based on an erroneous association with a particular species: verruculogen, viridicatumtoxin and rubratoxin come to mind. As time has gone on, and more and more compounds have been described, lists of species-mycotoxin associations have become so large, and the inaccuracies in them so widespread in acceptance, that determining true associations has become very difficult. It does not need to be emphasised how important it is that these associations be known accurately. The possible presence of mycotoxigenic fungi in foods, and rational decisions on the status of foods suspected to contain mycotoxins, are ever present problems in the food industry around the world. In defining mycotoxins, we exclude fungal metabolites which are active against bacteria, protozoa, and lower animals including insects. * J. C. Frisvad and U. Thrane, Center for Microbial Biotechnology, BioCentrumDTU, Technical University of Denmark, Building 221, DK-2800 Kgs. Lyngby, Denmark. Correspondence to [email protected] † R. A. Samson, Centraalbureau voor Schimmelcultures, PO Box 85167, 3508 AD Utrecht, Netherlands ‡ J. I. Pitt, Food Science Australia, CSIRO, PO Box 52, North Ryde, NSW 1670, Australia

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Jens C. Frisvad et al.

Furthermore we exclude Basidiomycete toxins, because these are ingested by eating fruiting bodies, a problem different from the ingestion of toxins produced by microfungi. The definition of microfungi is not rigorous, but understood here to refer principally to Ascomycetous fungi, including those with no sexual stage. Lower fungi, from the subkingdom Zygomycotina, i.e. genera such as Rhizopus and Mucor, are not excluded, but compounds of sufficient toxicity to be termed mycotoxins have not been found in these genera, except perhaps for rhizonin A and B from Rhizopus microsporus (Jennessen et al., 2005). This paper sets out to provide an up to date authoritative list of mycotoxins which are known to have caused, or we believe have the potential to cause, disease in humans or vertebrate animals, and the fungal species which have been shown to produce them. We believe that all of the important and known mycotoxins produced by Aspergillus, Fusarium and Penicillium species have been included in this list. However, it is possible that other species will be found which are capable of producing known toxins, or other toxins of consequence will arise. It is also important to note that there are many errors in the literature concerning the mycotoxins and the fungi which produce them (Frisvad et al., 2006). Many other toxic chemicals, known to be produced by species from these genera, have been excluded from this list for one reason or another. The very toxic chemicals, the janthitrems, have been excluded from this list because the species which make them, including P. janthinellum, normally do not grow to a significant extent in foods. On the other hand Penicillium tularense has recently been demonstrated to produce janthitrems in tomatoes (Andersen and Frisvad, 2004), so maybe these mycotoxins may occur sporadically. Other compounds which occur quite commonly in foods, including mycophenolic acid (Lafont et al., 1979, Lopez-Diaz et al., 1996; Overy and Frisvad, 2005), are of such low acute toxicity to vertebrate animals that their involvement in human or animal diseases appears unlikely. On the other hand mycophenolic acid has been reported to be strongly immunosuppressive (Bentley, 2000), so this fungal metabolite could pave the way for bacterial infections. Toxic low molecular weight compounds that may not be considered mycotoxins in a strict sense include aflatrem, botryodiploidin, brefeldin A, chetomin, chetocins, emestrin, emodin, engleromycin, fusarin C, lolitrems, paspalicine, paspaline, paspalinine, paspalitrems, paxilline, territrems, tryptoquivalins, tryptoquivalons, verruculotoxin, verticillins, and viridicatumtoxin which are among the fungal secondary metabolites listed as mycotoxins by Betina (1989).

Important Mycotoxins and the Fungi Which Produce Them

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Future research may show that some of these are more important for human and domestic animals health than currently indicated. For convenience the list below has been set out by genus, but it should be kept in mind that some mycotoxins are common to both Aspergillus and Penicillium species (Samson, 2001). The list below sets out to be encyclopaedic, but at the same time we have indicated, where possible, which species producing a particular toxin are more likely to occur in foods and which are probably of little consequence.

2.

ASPERGILLUS TOXINS

2.1.

Aflatoxins

Aflatoxins are potent carcinogens (Class 1; JECFA, 1997) affecting man and all tested animal species including birds and fish. Four compounds are commonly produced in foods: aflatoxins B1, B2, G1 and G2, named for the colour of their fluorescence under ultra violet light, and their relative position on TLC plates. Major sources. Aspergillus flavus is the most common species producing aflatoxins, occurring in most kinds of foods in tropical countries. This species has a special affinity with three crops, maize, peanuts and cottonseed, and usually produces only B aflatoxins. Only about 40% of known isolates produce aflatoxin. Aspergillus parasiticus occurs commonly in peanuts, but is quite rare in other foods. It is also restricted geographically, and is rare in Southeast Asia (Pitt et al., 1993). A. parasiticus produces both B and G aflatoxins, and virtually all known isolates are toxigenic. Minor sources. Table 1 shows the species which are known to be capable of producing aflatoxin in culture, and some details concerning their appearance and their occurrence. Note that most of the minor species are known from only a very few isolates, and their occurrence in foodstuffs or feedstuffs is at most rare. On the other hand A. nomius, A. toxicarius, and A. parvisclerotigenus may be more common than expected, because it is very difficult to distinguish between those species and isolates may easily have been identified as A. flavus or A. parasiticus.

2.2.

Cyclopiazonic acid (see also Penicillium)

Cyclopiazonic acid (CPA) (Holzapfel, 1968) is a potent mycotoxin that produces focal necrosis in most vertebrate inner organs in high

6 Jens C. Frisvad et al.

Table 1. Morphology and mycotoxin production characteristic of species in Aspergillus that are known aflatoxin producers a Species Heads Conidia Sclerotia Known occurrence Mycotoxins Aspergillus flavus Large, spherical Mostly biseriate Ubiquitous in Spherical to B aflatoxins ellipsoidal, smooth tropics and (40% of isolates); to finely rough walls subtropics CPAb, ca 50% A. parasiticus Rarely biseriate USA, South Spherical, rough walls B and G aflatoxins Large, spherical (uncommon) (nearly 100%) America, Australia A. nomius Mostly biseriate Small, bullet Spherical to USA, Thailand B and G aflatoxins ellipsoidal, smooth (usually) shaped to finely rough walls A. bombycis Not reported Mostly biseriate Japan, Indonesia Spherical to B and G aflatoxins (silkworms only) subspheroidal, roughened A. pseudotamarii Biseriate Spherical to Large, spherical Japan, Argentina B aflatoxins, CPA subspheroidal, very rough walls A. toxicarius Rarely biseriate USA, Uganda Sphaerical, rough B and G aflatoxins Large, sphaerical walled A. parvisclerotigenus Mostly biseriate USA, Argentina, Spherical, rough B and G aflatoxins, Small, sphaerical walled Japan, Nigeria CPA A. ochraceoroseus Biseriate Subspheroidal to B aflatoxins, Not reported Ivory Coast (soil) ellipsoidal, sterigmatocystin smooth walled A. rambellii Biseriate Ellipsoidal, smooth B aflatoxins, Not reported Ivory Coast (soil) walled sterigmatocystin Emericella astellata Biseriate Spheroidal, rugulose B aflatoxins, Ascomata and Ecuador (leaves walls hülle cells of Ilex) Sterigmatocystin Emericella B aflatoxins, Biseriate Sphaeroidal, Ascomata and Venezuela venezuelensis rugulose walls hülle cells (mangrove) Sterigmatocystin a From Kurtzman et al., 1987; Klich and Pitt, 1988; Pitt and Hocking, 1997; Klich et al., 2000; Ito et al., 2001; Peterson et al., 2001; Frisvad et al., 2004a; Frisvad and Samson, 2004a; Frisvad et al., 2005a. b CPA, cyclopiazonic acid.

Important Mycotoxins and the Fungi Which Produce Them

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concentrations and affects the ducts or organs originating from ducts. It was originally believed that aflatoxins were responsible for all the toxic effects of Aspergillus flavus contaminated peanuts to turkeys in Turkey X disease, but it was later shown that cyclopiazonic acid had an additional severe effect on the muscles and bones of the turkeys (Jand et al., 2005). Major sources. Aspergillus flavus and the domesticated form A. oryzae often produce large amounts of CPA. A. flavus is common on oil seeds, nuts, peanuts and cereals, but may also produce aflatoxin on dried fruits (Pitt and Hocking, 1997). Minor sources. Other producers of CPA in Aspergillus include A. tamarii, A. pseudotamarii, A. parvisclerotigenus, but the role of these fungi concerning CPA production in foods or feeds is not clear.

2.3.

Cytochalasin E

Cytochalasin E is a very toxic metabolite of Aspergillus clavatus. It may occur in malting barley (Lopez-Diaz and Flannigan, 1997) Major source. Aspergillus clavatus. Minor source. Rosellinia necatrix is not found in foods.

2.4.

Gliotoxin

Gliotoxin is strongly immunosuppressive, but is probably only a potential problem in animal feeds (Betina, 1989). Major sources. Aspergillus fumigatus has been found in animal feeds. Minor sources. Gliocladium virens, P. lilacinoechinulatum and few other soil-borne species also produce gliotoxin.

2.5.

β-Nitropropionic Acid (BNP)

β-nitropropionic acid has been reported to be involved in sugar cane poisoning of children, but may potentially also cause other intoxications, as producers are widespread (Burdock et al., 2001). Furthermore BNP has been found in miso, shoyu and katsuobushi and it can be produced by A. oryzae when artificially inoculated on cheese, peanuts etc. Unfortunately A. flavus has not been tested for the production of BNP, but BNP production by A. oryzae on peanuts indicates that A. flavus may be able to produce this mycotoxin in combination with aflatoxin B1, cyclopiazonic acid and kojic acid. The possible synergistic effect of these mycotoxins on mammals is unknown.

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Major sources. The BNP producing fungi from sugar cane are Arthrinium phaeospermum and Art. sacchari, but other species such as Art. terminalis, Art. saccharicola, Art. aureum and Art. sereanis also produce BNP (Burdock et al., 2001). A. flavus may be an important producer of this mycotoxin in foods, but there are no surveys that include analytical determination of BNP alongside cyclopiazonic acid and aflatoxin B1. A. oryzae and A. sojae can produce BNP in miso and shoyu, but it is probably more important that their wild-type forms, A. flavus and A. parasiticus respectively, may produce BNP in foods. More research is needed in this area. Minor sources. Penicillium atrovenetum is another authenticated producer of BNP, but this fungus is only found in soil. Incorrect sources. Penicillium cyclopium, P. chrysogenum, Aspergillus wentii, Eurotium spp., and A. candidus have been reported as producers of BNP (Burdock et al., 2001), but these identifications are doubtful.

2.6.

Ochratoxin A (see also Penicillium)

Ochratoxin A (OA) is a nephrotoxin, affecting all tested animal species, though effects in man have been difficult to establish unequivocally. It is listed as a probable human carcinogen (Class 2B) (JECFA, 2001). Links between OA and Balkan Endemic Nephropathy have long been sought, but not established (JECFA, 2001). Major sources. Aspergillus ochraceus (van der Merwe et al., 1965), occurring in stored cereals (Pitt and Hocking, 1997) and coffee (Taniwaki et al., 2003). A. ochraceus has been shown to consist of two species (Varga et al., 2000a, b; Frisvad et al., 2004b). The second and new species producing large amounts of ochratoxin A consistently, has been described as A. westerdijkiae. Actually the original producer of ochratoxin A from Andropogon sorghum in South Africa, NRRL 3174, has been designated as the type culture of A. westerdijkiae (Frisvad et al., 2004b). This is interesting as A. westerdijkiae is both a better and more consistent ochratoxin producer than A. ochraceus, and it may be also more prevalent in coffee than A. ochraceus. The ex type culture of A. ochraceus CBS 108.08 only produces trace amounts of ochratoxin A. Aspergillus carbonarius (Horie, 1995) is a major OA producer. It occurs in grapes, producing OA in grape products, including grape juice, wines and dried vine fruits (IARC, 2002; Leong et al., 2004) and sometimes on coffee beans (Taniwaki et al., 2003; Abarca et al., 2004). Aspergillus niger is an extremely common species, but only few strains

Important Mycotoxins and the Fungi Which Produce Them

9

appear to be producers of OA, so this species may be of much less importance than A. carbonarius in grapes, wine and green coffee beans (Abarca et al., 1994; Taniwaki et al., 2003; Leong et al., 2004). It may be of major importance, however, as A. niger NRRL 337, referred to as the “food fungus”, produces large amounts of OA in pure culture. This fungus is used for fermentation of potato peel waste etc. and used for animal feed (Schuster et al., 2002). Petromyces alliaceus (Lai et al., 1970), produces large amounts of ochratoxin A in pure culture, and OA produced by this fungus has been found in figs in California (Bayman et al., 2002). Aspergillus steynii, from the Aspergillus section Circumdati, is also a very efficient producer of OA, and has been found in green coffee beans, mouldy soy beans and rice (Frisvad et al., 2004b). As with A. westerdijkiae, A. steynii may have been identified as A. ochraceus earlier, so the relative abundance of these three species is difficult to evaluate at present. Penicillium verrucosum is the major producer of ochratoxin A in stored cereals (Frisvad, 1985; Pitt, 1987; Lund and Frisvad, 2003). Penicillium nordicum (Larsen et al., 2001) is the main OA producer found in meat products such as salami and ham. Both OA producing Penicillium species have been found on cheese also, but have only been reported to be of high occurrence on Swiss hard cheeses (as P. casei, Staub, 1911). The ex type culture of P. casei is a P. verrucosum (Larsen et al., 2001). Minor sources. Several Aspergilli can produce ochratoxin A in large amounts, but they appear to be relatively rare. In Aspergillus section Circumdati (formerly the Aspergillus ochraceus group), the following species can produce ochratoxin A: Aspergillus cretensis, A. flocculosus, A. pseudoelegans, A. roseoglobulosus, A. sclerotiorum, A. sulphureus and Neopetromyces muricatus (Frisvad et al., 2004b). According to Ciegler (1972) and Hesseltine et al. (1972) A. melleus, A. ostianus, A. persii and A. petrakii may produce trace amounts of OA, but this has not been confirmed since publication of those papers. Strains of these species reported to produce large amounts of OA were reidentified by Frisvad et al. (2004b). In Aspergillus section Flavi, Petromyces albertensis produces ochratoxin A. In Aspergillus section Nigri, A. lacticoffeatus and A. sclerotioniger produce ochratoxin A (Samson et al., 2004).

2.7.

Sterigmatocystin

Sterigmatocystin is a possible carcinogen. However, its low solubility in water or gastric juices limits its potential to cause human illness (Pitt and Hocking, 1997).

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Major sources. The major source of sterigmatocystin in foods is Aspergillus versicolor. This fungus is common on cheese, but may also occur on other substrates (Pitt and Hocking, 1997). Minor sources. A large number of species are able to produce sterigmatocystin, including Chaetomium spp., Emericella spp., Monocillium nordinii and Humicola fuscoatra (Joshi et al., 2002). These species are unlikely to contaminate foods.

2.8.

Verruculogen and Fumitremorgins

Verrucologen is an extremely toxic tremorgenic mycotoxin, but it is unlikely to be founding significant levels in foods. Neosartorya fisheri may be present in heat treated foods, but N. glabra and allied species are much more common in foods, and the latter species do not produce verrucologen. Major sources. Aspergillus fumigatus and Neosartorya fischeri are the major Aspergillus species producing verruculogen but these species are uncommon in foods. These species produce many other toxic compounds including gliotoxin, fumigaclavins, and tryptoquivalins (Cole et al., 1977; Cole and Cox, 1981; Panaccione and Coyle, 2005). Minor sources. Aspergillus caespitosus, Penicillium mononematosum and P. brasilianum are efficient producers of verrucologen and fumitremorgins, but are very rare in foods and feeds.

3.

FUSARIUM TOXINS

3.1.

Antibiotic Y

Antibiotic Y has significant antibiotic properties towards phytopathogenic bacteria but low cell toxicity (Golinski et al., 1986). However, this compound, which originally was named lateropyrone (Bushnell et al., 1984), has not been studied in detail. Producers of antibiotic Y are widespread and common in agricultural products, so the natural occurrence of antibiotic Y may be of importance. Natural occurrence in cherries, apples and wheat grains has been reported (Andersen and Thrane, 2005). Major sources. The main producer is Fusarium avenaceum which occurs frequently in cereal grain, fruit and vegetables. Another consistent producer is F. tricinctum, which also is very frequently found on cereal grains in temperate climates.

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Minor sources. F. lateritium is known as a plant pathogen, but also causes spoilage in fruits and has been reported from apples and cherries in which antibiotic Y was detected (Andersen and Thrane, 2006). In warmer climates F. chlamydosporum is a potential producer of antibiotic Y in cereal grain and other seeds.

3.2.

Butenolide

Butenolide is a collective name for compounds with a given ring structure; however in Fusarium mycotoxicology butenolide is a synonym for 4-acetamido-2-buten-4-olide, which has been associated with cattle diseases (fescue foot) since the mid 1960s (Yates et al., 1969). The toxicology has been thoroughly discussed by Marasas et al. (1984). There have been no reports of butenolide in foods, but it may be an important toxin due to the reported synergistic effect with enniatin B (Hershenhorn et al., 1992). Major sources. The original reported producer of butenolide is F. sporotrichioides [reported as F. nivale, see Marasas et al. (1984) for details] and other frequent producers of butenolide in cereals are F. graminearum and F. culmorum. Minor sources. Other potential producers of butenolide are F. avenaceum, F. poae and F. tricinctum which are frequently found in cereal grains together with F. crookwellense, F. sambucinum and F. venenatum. The latter three species also can be found in potatoes and other root vegetables.

3.3.

Culmorin

Culmorin has a low toxicity in several biological assays (Pedersen and Miller, 1999) but a synergistic effect with deoxynivalenol towards caterpillars has been demonstrated (Dowd et al., 1989). Culmorin and hydroxyculmorins have been detected in cereals (Ghebremeskel and Langseth, 2000). These samples also contained deoxynivalenol and acetyl-deoxynivalenol. Major sources. F. culmorum and F. graminearum, found in cereals, are the major producers of culmorin. The less widely distributed species F. poae and F. langsethiae are also consistent producers of culmorin and derivatives (Thrane et al., 2004). Minor sources. Other species producing culmorin are F. crookwellense and F. sporotrichioides, also found in cereals.

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Cyclic Peptides

The two groups of cyclic peptides, beauvericin and enniatins, are structurally related and they show antibiotic and ionophoric activities (Kamyar et al., 2004). Both groups of cyclic peptides have been detected in agricultural products (Jestoi et al., 2004). 3.4.1.

Beauvericin

Beauvericin was originally found in entomopathogenic fungi such as Beauveria bassiana and Isaria fumosorosea (formerly Paecilomyces fumosoroseus; Luangsa-Ard et al., 2005) but has also been detected in several Fusarium species occurring on food (Logrieco et al., 1998). Major sources. Fusarium subglutinans, F. proliferatum and F. oxysporum are consistent producers of beauvericin and have often been found to produce high quantities under laboratory conditions. These species are often found on maize and fruits. Minor sources. Several species of the Gibberella fujikuroi complex have been reported to produce beauvericin in low amounts, including F. nygamai, F. dlaminii and F. verticillioides from cereals and fruits. The systematics of these Fusaria has developed dramatically during the last years, so a lot of species specific information of toxin production is not available. F. avenaceum, F. poae and F. sporotrichioides on cereal grain, fruits and vegetables are known to produce beauvericin in low amounts (Morrison et al., 2002; Thrane et al., 2004). In addition, F. sambucinum and a few strains of F. acuminatum, F.equiseti and F. longipes from agricultural products have also been reported low producers of beauvericin (Logrieco et al., 1998). 3.4.2.

Enniatins

Enniatins are a group of more than 15 related compounds produced by several Fusarium species, but also from Halosarpeia sp. and Verticillium hemipterigenum; however these are not of food origin. Major sources. Fusarium avenaceum is the most important enniatin producer in cereals and other agricultural food plants, because this species is a very frequent and consistent producer of enniatin B (Morrison et al., 2002). Fusarium sambucinum is a consistent producer of enniatin B and diacetoxyscirpenol and causes dry rot in potatoes; however the role of these toxins has not been examined.

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Minor sources. F. langsethiae, F. poae and F. sporotrichioides, mainly occur on cereal grain, F. lateritium from fruits and F. acuminatum from herbs.

3.5.

Fumonisins

Since the discovery of fumonisins in the late 1980s much attention has been paid to these highly toxic compounds. Several reviews on the chemistry, toxicology and mycology have been published (Marasas et al., 2001; Weidenbörner, 2001). Major sources. F. verticillioides (formerly known as F. moniliforme; Seifert et al., 2003) and F. proliferatum are the main sources of fumonisins in maize. These species and fumonisins in maize and to a lesser extent other cereal crops have been reported from all over the world in numerous papers and book chapters. Minor sources. Other fumonisin producing species are Fusarium nygamai, F. napiforme, F. thapsinum, F. anthophilum and F. dlamini from millet, sorghum and rice. Some strains of these species have also been isolated from soil debris.

3.6.

Fusaproliferin

Fusaproliferin is a recent discovered mycotoxin which shows teratogenic and pathological effects in cell assays (Bryden et al., 2001). Fusaproliferin has been detected in natural samples together with beauvericin and fumonisin (Munkvold et al., 1998). Nothing is known about a possible synergistic effect in such toxin combinations. Major sources. Fusarium proliferatum and F. subglutinans are the major sources in maize and other cereal grains. The fungi and fusaproliferin have been detected in Europe, North America and South Africa (Wu et al., 2003). Minor sources. A few strains of F. globosum, F. guttiforme, F. pseudocircinatum, F. pseudonygamai and F. verticillioides have been found to produce fusaproliferin, however the systematics in this section of Fusarium has developed dramatically within recent years so specific information on the toxin production by recently described species is unknown.

3.7.

Moniliformin

Moniliformin is cytotoxic, inhibits protein synthesis and enzymes, causes chromosome damages and induces heart failure in mammals

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and poultry (Bryden et al., 2001). Moniliformin has been found world wide in cereal samples Major sources. In maize F. proliferatum and F. subglutinans are the main producers of moniliformin, whereas F. avenaceum and F. tricinctum are the key sources in cereals grown in temperate climates. Minor sources. In sorghum, millet and rice F. napiforme, F. nygamai, F. verticillioides and F. thapsinum may be responsible for moniliformin production. Some strains of F. oxysporum produce a significant amount of moniliformin under laboratory condition; however there is no detailed information on a possible production in vegetables and fruits. An overview of other minor sources has been published (Schütt et al., 1998).

3.8.

Trichothecenes

More than 200 trichothecenes have been identified and the nonmacrocyclic trichothecenes are among the most important mycotoxins. Trichothecenes are haematotoxic and immunosuppressive. In animals, vomiting, feed refusal and diarrhoea are typical symptoms. Skin oedema in humans has also been observed. An EU working group on has reported on trichothecenes in food (Schothorst and van Egmond, 2004). 3.8.1.

Deoxynivalenol (DON) and Acetylated Derivatives (3ADON, 15ADON)

Deoxynivalenol (DON) and its acetylated derivatives (3ADON, 15ADON) are by far the most important trichothecenes. Numerous reports on world-wide occurrence have been published and several international symposia and workshops have focussed on DON (Larsen et al., 2004). Major sources. Fusarium graminearum and F. culmorum are consistent producers of DON, especially in cereals. Within both species strains have been grouped into those that produce DON and its derivatives, and those that produce nivalenol and furarenon X as their major metabolites. Intermediates have also been found (Nielsen and Thrane, 2001). Recently, F. graminearum has been divided into nine phylogenetic species (O’Donnell et al., 2004); however in the present context this species concept will not be used as a correlation to existing mycotoxicological literature is impossible at this stage. Minor sources. Production of DON by F. pseudograminearum has been reported, but this species is restricted to warmer climates.

Important Mycotoxins and the Fungi Which Produce Them

3.8.2.

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Nivalenol (NIV) and Fusarenon X (FX, 4ANIV)

Nivalenol (NIV) and fusarenon X (FX, 4ANIV) occur in the same commodities as DON and are in many cases covered by the same surveys due to the high degree of similiarity. NIV is often detected in much lower concentrations than DON, but is considered to be more toxic. Major sources. Fusarium graminearum is a well known producer of NIV and FX in cereals. In temperate climates F. poae, which is a consistent producer of NIV (Thrane et al., 2004), may be responsible for NIV in cereals. Minor sources. Strains of F. culmorum that produce NIV are less commonly isolated than those that produce DON producers. F. equiseti and F. crookwellense found in some cereal samples and in vegetables may also produce NIV. In potatoes F. venenatum strains that produce NIV have been detected (Nielsen and Thrane, 2001). 3.8.3.

T-2 toxin

T-2 toxin is one of the most toxic trichothecenes, whereas the derivative HT-2 toxin is less toxic. Due to structural similarity these toxins are often included in the same analytical method. Major sources. Fusarium sporotrichioides and F. langsethiae, frequently isolated from cereals in Europe, are consistent producers of T-2 and HT-2 (Thrane et al., 2004). Minor sources. Only a few T-2 and HT-2 producing strains of F. poae and F. sambucinum have been found (Nielsen and Thrane, 2001; Thrane et al., 2004). 3.8.4.

Diacetoxyscirpenol (DAS)

Diacetoxyscirpenol (DAS) and monoacetylated derivatives (MAS) are a fourth group of important trichothecenes in food. Major sources. Fusarium venenatum isolates often produce high levels of DAS and this species is frequently isolated from cereals and potatoes (Nielsen and Thrane, 2001). F. poae isolates also often produce high levels of DAS. Minor sources. Fusarium equiseti isolates can produce DAS and MAS in high amounts, but this species is infrequently isolated from cereals and vegetables. F. sporotrichioides and F. langsethiae also produce DAS and MAS; however at lower levels (Thrane et al., 2004). F. sambucinum isolates produce DAS and MAS and are a probable cause of DAS in potatoes (Ellner, 2002).

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Zearalenone

Zearalenone causes hyperoestrogenism in swine and possible effects in humans have also been reported. Derivatives of zearalenone have been used as growth promoters in livestock; however this is now banned in European Union (Launay et al., 2004). The toxicity of zearalenone and its derivatives have been reviewed recently (Hagler et al., 2001). Major sources. Fusarium graminearum and F. culmorum are the most pronounced producers of zearalenone and several derivatives. They occur frequently in cereals all over the world. Recently, F. graminearum has been divided into nine phylogenetic species (O’Donnell et al., 2004); however in the present context this species concept will not be used as a correlation to existing mycotoxicological literature is impossible at this stage. Minor sources. Under laboratory conditions Fusarium equiseti produces a number of zearalenone derivatives in high amounts, but little is known about production under natural conditions. F. crookwellense also produces zearalenone.

4.

PENICILLIUM TOXINS

4.1.

Chaetoglobosins

The chaetoglobosins are toxic compounds that may be involved in mycotoxicosis. They are produced by common food-borne Penicillia and have been found to occur naturally (Andersen et al., 2004). Major sources. Penicillium expansum and P. discolor are major sources of the chaetoglobosins. Both species cause spoilage in fruits and vegetables, and the latter species also occurs on cheese (Frisvad and Samson, 2004b). Minor sources. Chaetomium globosum and P. marinum are probably not of significance in foods.

4.2.

Citreoviridin

Citreoviridin was reported as a cause of acute cardiac beriberi (Ueno, 1974), but a more in depth toxicological evaluation of this metabolite is needed. It has been associated with yellow rice disease,

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but this disease has also been associated with P. islandicum and its toxic metabolites cyclic peptides cyclochlorotine and islanditoxin, and anthraquinones luteoskyrin and rugulosin (Enomoto and Ueno, 1974). Major sources. Eupenicillium cinnamopurpureum has been found in cereals in USA and in Slovakia (Labuda and Tancinova, 2003) and is an efficient producer of citreoviridin. P. citreonigrum may be of some importance in yellowed rice. Minor sources. P. smithii, P. miczynskii and P. manginii (Frisvad and Filtenborg, 1990) have most often been recovered from soil and only rarely from foods. Aspergillus terreus has occasionally been reported from foods, but is primarily a soil-borne fungus.

4.3.

Citrinin

Citrinin is a nephrotoxin, but probably of less importance than ochratoxin A (Reddy and Berndt, 1991), however, producers of citrinin are widespread and common in foods. Citrinin has been found in cereals, peanuts and meat products (Reddy and Berndt, 1991). Major sources. P. citrinum is an efficient and consistent producer of citrinin and has been found in foods world-wide (Pitt and Hocking, 1997). P. verrucosum is predominantly cereal-borne in Europe and often produces citrinin as well as ochratoxin A (Frisvad et al., 2005b). P. expansum, common in fruits and other foods, sometimes produces citrinin. P. radicicola is commonly found in onions, carrots and potatoes (Overy and Frisvad, 2003). Minor sources. Aspergillus terreus, A. carneus, P. odoratum and P. westlingii have been reported as producers of citrinin, but are not likely to occur often in foods.

4.4.

Cyclopiazonic acid (see also Aspergillus)

Major sources. Penicillium commune and its domesticated form P. camemberti, and the closely related species P. palitans, are common on cheese and meat products and may produce cyclopiazonic acid in these products (Frisvad et al., 2004c). P. griseofulvum is also a major producer of cyclopiazonic acid, and may occur in long stored cereals and cereal products such as pasta (Pitt and Hocking, 1997). Minor sources. P. dipodomyicola occurs in the environs of the kangaroo rat in the USA, but has also been reported from rice in Australia and in a chicken feed mixture in Slovakia (Frisvad and Samson, 2004b).

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Mycophenolic acid

Despite having a low acute toxicity, mycophenolic acid may be a very important indirect mycotoxin as it highly immunosuppressive, perhaps influencing the course of bacterial and fungal infections (Bentley, 2000). Major sources. Penicillium brevicompactum is a ubiquitous species and may produce mycophenolic acid in foods, e.g. ginger (Overy and Frisvad, 2005). Two other major species producing mycophenolic acid are P. roqueforti and P. carneum. Another important producer is Byssochlamys nivea (Puel et al., 2005). Mycophenolic acid has been found to occur naturally in blue cheeses (Lafont et al., 1979). Minor sources. The soil-borne species Penicillium fagi also produces mycophenolic acid (Frisvad and Filtenborg, 1990, as P. raciborskii). Septoria nodorum (Devys et al., 1980) is another source but is unimportant as a food contaminant.

4.6.

Ochratoxin A (see also Aspergillus)

Major sources. Penicillium verrucosum (Frisvad, 1985; Pitt, 1987) is the major producer of ochratoxin A in cool climate stored cereals (Lund and Frisvad, 2003). Penicillium nordicum (Larsen et al., 2001) is the main OA producer found in manufactured meat products such as salami and ham. Both OA producing Penicillium species have been found on cheese also, but have only been reported to be of high occurrence on Swiss hard cheeses (as P. casei Staub, 1911). The ex type culture of P. casei is a P. verrucosum (Larsen et al., 2001).

4.7.

Patulin

Patulin is generally very toxic for both prokaryotes and eukaryotes, but the toxicity for humans has not been conclusively demonstrated. Several countries in Europe and the USA have now set limits on the level of patulin in apple juice. Major sources. Penicillium expansum is by far the most important source of patulin. P. expansum is the major species causing spoilage of apples and pears, and is the major source of patulin in apple juice and other apple and pear products. Byssochlamys nivea may be present in pasteurised fruit juices and may produce patulin and mycophenolic acid (Puel et al., 2005).

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Penicillium griseofulvum is a very efficient producer of high levels of patulin in pure culture, and it may potentially produce patulin in cereals, pasta and similar products. P. carneum may produce patulin in beer, wine, meat products and rye-bread as it has been found in those substrates (Frisvad and Samson, 2004b), but there are no reports yet on patulin production by this species in those foods. P. carneum also produces mycophenolic acid, roquefortine C and penitrem A (Frisvad et al., 2004c). P. paneum occurs in rye-bread (Frisvad and Samson, 2004b), but again actual production of patulin in this product has not been reported. P. sclerotigenum is common in yams and has the ability to produce patulin in laboratory cultures. Minor sources. The coprophilous fungi P. concentricum, P. clavigerum, P. coprobium, P. formosanum, P. glandicola, P. vulpinum, Aspergillus clavatus, A. longivesica and A. giganteus are very efficient producers of patulin in the laboratory, but only A. clavatus may play any role in human health, as it may be present in beer malt (LopezDiaz and Flannigan, 1997). Aspergillus terreus, Penicillium novae-zeelandiae, P. marinum, P. melinii and other soil-borne fungi may produce patulin in pure culture, but are less likely to occur in any foods.

4.8.

Penicillic acid

Penicillic acid (Alsberg and Black, 1911) and dehydropenicillic acid (Obana et al., 1995) are small toxic polyketides, but their major role in mycotoxicology may be in their possible synergistic toxic effect with OA (Lindenfelser at al., 1973; Stoev et al., 2001) and possible additive or synergistic effect with the naphtoquinones hepatotoxins xanthomegnin, viomellein and vioxanthin. Major sources. Penicillic acid is likely to co-occur with OA, xanthomegnin, viomellein and vioxanthin produced by members of Aspergillus section Circumdati and Penicillium series Viridicata (which often co-occur with P. verrucosum). The Aspergillus species often occur in coffee and the Penicillia are common in cereals. The major sources of penicillic acid are P. aurantiogriseum, P. cyclopium, P. melanoconidium and P. polonicum (Frisvad and Samson, 2004b) and all members of Aspergillus section Circumdati (Frisvad and Samson, 2000). Penicillic acid is produced by P. tulipae and P. radicicola, which are occasionally found on onions, carrots and potatoes (Overy and Frisvad, 2003). Minor sources. Penicillic acid has been found in one strain of P. carneum (Frisvad and Samson, 2004b).

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Penitrem A

Penitrem A is a highly toxic tremorgenic indol-terpene. It has primarily been implicated in animal mycotoxicoses (Rundberget and Wilkins, 2002), but has also been suspected to cause tremors in humans (Cole et al., 1983; Lewis et al., 2005). Major sources. Penicillium crustosum is the most important producer of penitrem A (Pitt, 1979). This species is of world-wide distribution and often found in foods. This mycotoxins is produced by all isolates of P. crustosum examined (Pitt, 1979; Sonjak et al., 2005). P. melanoconidium is common in cereals (Frisvad and Samson, 2004b), but it is not known whether this species can produce penitrem A in infected cereals. Minor sources. P. glandicola, P. clavigerum, and P. janczewskii are further producers of penitrem A (Ciegler and Pitt, 1970; Frisvad and Samson, 2004b; Frisvad and Filtenborg, 1990), but have been recovered from foods only sporadically.

4.10.

PR toxin

PR toxin is a mycotoxin that is acutely toxic and can damage DNA and proteins (Moule et al., 1980; Arnold et al., 1987). It is unstable in cheese (Teuber and Engel, 1983), but it may be produced in silage and other substrates. Major sources. Penicillium roqueforti is the major source of PR toxin. It has been reported also from P. chrysogenum (Frisvad and Samson, 2004b).

4.11.

Roquefortine C

The status of roquefortine C as a mycotoxin has been questioned, but it is a very widespread fungal metabolite, and is produced by a large number of species. The acute toxicity of roquefortine C is not very high (Cole and Cox, 1981), but it has been reported as a neurotoxin. Major sources. Penicillium albocoremium, P. atramentosum, P. allii, P. carneum, P. chrysogenum, P. crustosum, P. expansum, P. griseofulvum, P. hirsutum, P. hordei, P. melanoconidium, P. paneum, P. radicicola, P. roqueforti, P. sclerotigenum, P. tulipae and P. venetum are all producers that have been found in foods, but the natural occurrence of roquefortine C has been reported only rarely.

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Minor sources. P. concentricum, P. confertum, P. coprobium, P. coprophilum, P. flavigenum, P. glandicola, P. marinum, P. persicinum and P. vulpinum are less likely food contaminants.

4.12.

Rubratoxin

Rubratoxin is a potent hepatotoxin (Engelhardt and Carlton, 1991) and is of particular interest as it has been implicated in severe liver damage in three Canadian boys, who drank rhubarb wine contaminated with Penicillium crateriforme. One of the boys needed to have the liver transplanted (Richer et al., 1997). Major producers. P. crateriforme is the only known major producer of rubratoxin A and B (Frisvad, 1989).

4.13.

Secalonic Acid D

The toxicological data on secalonic acid D and F are somewhat equivocal (Reddy and Reddy, 1991), so the significance of this metabolite in human and animal health is somewhat uncertain. Major sources. Claviceps purpurea, Penicillium oxalicum, Phoma terrestris and Aspergillus aculeatus produce large amounts of secalonic acid D and F in pure culture. Secalonic acid D has been found to occur in grain dust in USA (Palmgren, 1985; Reddy and Reddy, 1991).

4.14.

Verrucosidin

Verrucosidin is a of the mycotoxin from species in Penicillium series Viridicata that has been claimed to cause mycotoxicosis in animals (Burka et al., 1983). Major sources. Penicillium polonicum, P. aurantiogriseum and P. melanoconidium are the major known sources of verrucosidin (Frisvad and Samson, 2004b).

4.15.

Xanthomegnin, Viomellein and Vioxanthin

These toxins have been reported to cause experimental mycotoxicosis in pigs and they apparently are more toxic to the liver than to kidneys in mammals (Zimmerman et al., 1979). They have been found to be naturally occurring in cereals (Hald et al., 1983; Scudamore et al., 1986).

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Major sources. P. cyclopium, P. freii, P. melanoconidium, P. tricolor and P. viridicatum are common in cereals. A. ochraceus, A. westerdijkiae and possibly A. steynii are common in green coffee beans and are occasionally found in grapes and on rice. Minor sources. P. janthinellum and P. mariaecrucis are soil-borne species producing these hepatotoxins (Frisvad and Filtenborg, 1990).

5.

TOXINS FROM OTHER GENERA

5.1.

Claviceps Toxins

Ergot alkaloids are common in sclerotia of Claviceps, which are produced on cereals, especially in whole rye. These sclerotia are often removed before milling of the rye, and outbreaks of ergotism rarely occur now. Major sources. Claviceps purpurea and C. paspali are the major sources of ergot alkaloids (Blum, 1995). Several Penicillia and Aspergilli can produce clavinet type alkaloids also, but their possible role in mycotoxicology is unknown.

5.2.

Alternaria Toxins

Tenuazonic acid is regarded as the most toxic of the secondary metabolites from Alternaria (Blaney, 1991). It is also produced by a Phoma species. Major sources. Phoma sorghina appears to be the most important producer of tenuazonic acid. It has been associated with onyalai, a haematological disease (Steyn and Rabie, 1976). Species in the Alternaria tenuissima complex often produce tenuazonic acid, but it has not been found in isolates of A. alternata sensu stricto. A. citri, A. japonica, A. kikuchiana, A. longipes, A. mali, A. oryzae, and A. solani have also been reported to produce tenuazonic acid (Sivanesan, 1991). Many other metabolites have been found in Alternaria, and some can occur naturally in tomatoes, apples and other fruits (Sivanesan, 1991; Andersen and Frisvad, 2004). The toxicity of such compounds, including alternariols, is not well examined.

5.3.

Phoma and Phomopsis Toxins

Lupinosis toxin (phomopsin) is produced when Phomopsis leptostromiformis grows on lupin plants (Lupinus species) and lupin seeds

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23

(Culvenor et al., 1977). It is a hepatotoxin which has caused widespread disease in sheep grazing lupins in Australia, South Africa and parts of Europe (Marasas, 1974; Culvenor et al., 1977). As lupin seed is used for human food in South Asia, quality control of phomopsin is important.

5.4.

Pithomyces Toxins

Sporidesmin is produced by Pithomyces chartarum and causes facial eczema in sheep (Atherton et al., 1974). However, this is a disease of pasture only.

5.5.

Stachybotrys Toxins

Stachybotrys and Memnoniella spp. are primarily of importance for indoor air, but stachybotrytoxicosis was one of the first equine mycotoxicosis to be reported (Rodrick and Eppley, 1974). Stachybotrys chartarum and S. chlorohalonata are the two important fungi producing cyclic trichothecenes (satratoxins) and toxic atranones (Andersen et al., 2003; Jarvis, 2003).

5.6.

Monascus Toxins

Monascus ruber is used in the production of red rice in the Orient, and is a source of red food colouring. However, it has been repeatedly reported to produce citrinin (Blanc et al., 1995).

6.

DISCUSSION

A large number of filamentous fungi are able to produce secondary metabolites that are toxic to vertebrate animals, i.e. mycotoxins. Only a fraction of these fungi can produce mycotoxins in food or feeds, and among those, pathogenic field fungi and deteriorating storage fungi are the most significant. When misidentified fungi are excluded, only a few fungal species are highly toxigenic, and producing their toxins in sufficiently large amounts to cause public alarm. The most important among these are trichothecenes, fumonisins, aflatoxin, ochratoxin A and zearalenone (Miller, 1995), because their fungal producers are widespread and can grow and produce their toxins on many kinds of foods. Other mycotoxins are important, but may

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only occur on a single type of crop and cause mycotoxicosis in one kind of animal. Phomopsin is an example of this. Correct identification and knowledge of the associated mycobiota of the different foods and feeds will assist in determining which mycotoxins to look for. There have been examples of mycotoxins analysis for aflatoxin, trichothecenes, zearalenone, fumonisin and ochratoxin A in silage, where the dominant mycobiota is P. roqueforti, P. paneum, Monascus ruber and Byssochlamys nivea. In that particular case patulin, mycophenolic acid, PR toxin, and citrinin would be more relevant mycotoxins to analyse for. Rapid methods may are effective to secure healthy foods and feeds, but such methods should be based on mycological and ecological knowledge. We hope that our compilation of mycotoxin producers will help in deciding the most appropriate mycotoxin analyses of foods and feeds. New mycotoxins and new mycotoxin producers will no doubt appear, but we believe that the most important ones are listed here.

7.

ACKNOWLEDGEMENTS

Jens Frisvad and Ulf Thrane thank LMC and the Technical Research Council for financial support.

8.

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Pitt, J. I., 1987, Penicillium viridicatum, P. verrucosum, and the production of ochratoxin A, Appl. Environ. Microbiol. 53:266-269. Pitt, J. I., and Hocking, A. D., 1997, Fungi and Food Spoilage, 2nd edition, Blackie Academic and Professional, London, 596 pp. Pitt, J. I., Hocking, A. D., Bhudhasamai, K., Miscamble, B. F., Wheeler, K. A., and Tanboon-Ek, P., 1993, The normal mycoflora of commodities from Thailand. 1. Nuts and oilseeds, Int. J. Food Microbiol. 20:211-226. Puel, O., Tadrist, S. Galtier, P., Oswald, I. P., and Delaforge, M., 2005, Byssochlamys nivea as a source of mycophenolic acid, Appl. Environ. Microbiol. 71:550-553. Reddy, R.,V., and Berndt, W. O., 1991, Citrinin, in: Mycotoxins and Phytoalexins, R. P. Sharma and D. K. Salunkhe, eds, CRC Press, Boca Raton, Florida., pp. 237-250. Reddy, C. S. and Reddy, R. V. 1991. Secalonic acids, in: Mycotoxins and Phytoalexins, R. P. Sharma and D. K. Salunkhe, eds, CRC Press, Boca Raton, Florida., pp. 167190. Richer, L., Sigalet, D., Kneteman, N., Jones, A., Scott, R. B., Ashbourne, R., Sigler, L., Frisvad, J., and Smith, L., 1997, Fulminant hepatic failure following ingestion of moldy homemade rhubarb wine, Gastenterology 112: A1366. Rodrick, J. V., and Eppley, R. M., 1974, Stachybotrys and stachybotryotoxicosis, in: Mycotoxins, I. F. H. Purchase, ed., Elsevier, Amsterdam, pp. 181-197. Rundberget, T., and Wilkins, A. L., 2002, Thomitrems A and E, two indole-alkaloid isoprenoids from Penicillium crustosum Thom, Phytochemistry 61:979-985. Samson, R. A., 2001, Current fungal taxonomy and mycotoxins, in: Mycotoxins and phycotoxins in perspective at the turn of the century, de Koe, W. J., Samson, R. A., van Egmond, H. P., Gilbert, J., and Sabino, M., eds, Proceedings of the X international IUPAC Symposium, Mycotoxins and Phycotoxins, Guaruja-Sao Paulo, Brazil-May 21-25, 2000, W. J. de Koe, Wageningen, pp. 343-350. Samson, R. A., Houbraken, J. A. M. P., Kuijpers, A. F. A., Frank, J. M., and Frisvad, J. C., 2004, New ochratoxin A or sclerotium producing species in Aspergillus section Nigri, Stud. Mycol. (Utrecht) 50:45-61. Schothorst, R. C., and van Egmond, H. P., 2004, Report from SCOOP task 3.2.10, Collection of occurrence data of Fusarium toxins in food and assessment of dietary intake by the population of EU member states; Subtask: trichothecenes, Toxicol. Lett. 153:133-143. Schuster, E., Dunn-Coleman, N., Frisvad, J. C., and van Dijck, P. W. M., 2002, On the safety of Aspergillus niger -a review, Appl. Microbiol. Biotechnol. 59:426-435. Schütt, F., Nirenberg, H. I., and Deml, G., 1998, Moniliformin production in the genus Fusarium, Mycotoxin Res. 14:35-40. Scudamore, K. A., Atkin, P., and Buckle, A. E., 1986, Natural occurrence of the naphthoquinone mycotoxins, xanthomegnin, viomellein and vioxanthin in cereals and animal foodstuffs, J. Stored Prod. Res. 22:81-84. Seifert, K. A., Aoki, T., Baayen, R. P., Brayford, D., Burgess, L. W., Chulze, S., Gams, W., Geiser, D., de Gruyter, J., Leslie, J. F., Logrieco, A., Marasas, W. F. O., Nirenberg, H. I., O’Donnell, K., Rheeder, J. P., Samuels, G. J., Summerell, B. A., Thrane, U. and Waalwijk, C., 2003, The name Fusarium moniliforme should no longer be used, Mycol. Res. 107:643-644. Sivanesan, A., 1991, The taxonomy and biology of dematiaceous hyphomycetes and their mycotoxins, in: Fungi and mycotoxins in stored products, B. R. Champ, E. Highley, A. D. Hocking and J. I. Pitt, eds, ACIAR Proceedings No. 36, Australian Centre for International Agricultural Research, Canberra, pp. 47-64.

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Sonjak, S., Frisvad, J. C. and Gunde-Cimerman, N., 2005, Comparison of secondary metabolite production by Penicillium crustosum strains, isolated from Arctic and other various ecological niches, FEMS Microbiol. Ecol. 53: 51-60. Staub, W., 1911, Penicillium casei n. sp. als Ursache die rotbraunen Rinderfarbung bei Emmenthaler Käsen, Centrabl. f. Bakt. (II) 31:454. Steyn, P. S., and Rabie, C. J., 1976, Characterisation of magnesium and calcium tenuazonate from Phoma sorghina, Phytochemistry 15:1977-1979. Stoev, S. D., Vitanov, S., Anguelov, G., Petkova-Bocharova, T., and Creppy, E. E., 2001, Experimental mycotoxic nephropathy in pigs provoked by a diet containing ochratoxin A and penicillic acid, Vet. Res. Commun. 25:205-223. Taniwaki, M. H., Pitt, J. I., Teixeira, A. A., and Iamanaka, B. T., 2003, The source of ochratoxin A in Brazilian coffee and its formation in relation to processing methods, Int. J. Food Microbiol. 82:173-179. Teuber, M., and Engel, G., 1983, Low risk of mycotoxin production in cheese, Microbiol. Alim. Nutr. 1:193-197. Thrane, U., Adler, A., Clasen, P.-E., Galvano, F., Langseth, W., Lew, H., Logrieco, A., Nielsen, K. F., and Ritieni, A., 2004, Diversity in metabolite production by Fusarium langsethiae, Fusarium poae, and Fusarium sporotrichioides, Int. J. Food Microbiol. 95:257-266. Ueno, Y., 1974, Citreoviridin from Penicillium citreo-viride Biourge, in: Mycotoxins I. F. H. Purchase, ed., Elsevier, Amsterdam, pp. 283-302. Van der Merwe, K. J., Steyn, P. S., Fourie, L., Scott., D. B., and Theron, J. J., 1965, Ochratoxin A, a toxic metabolite produced by Aspergillus ochraceus Wilh., Nature 205:1112-1113. Varga, J., Tóth, B., Rigó, K, Téren, J, Hoekstra, R. F., and Kozakiewics, Z., 2000a, Phylogenetic analysis of Aspergillus section Circumdati based on sequences of the internal transcribed spacer regions of the 5.8 S rRNA gene, Fungal Gen. Biol. 30:71-80. Varga, J., Kevei, É., Tóth, B., Kozakiewicz, Z., and Hoekstra, R. F., 2000b, Molecular analysis of variability within the toxigenic Aspergillus ochraceus species, Can. J. Microbiol. 46:593-599. Weidenbörner, M., 2001, Food and fumonisins, Eur. Food Res. Technol. 212:262-273. Wu, X., Leslie, J. F., Thakur, R. A., and Smith, J. S., 2003, Purifiction of fusaproliferin form cultures of Fusarium subglutinans by preparative high-performance liquid chromatography, J. Agric. Food Chem. 51: 383-388. Yates, S. G., Tookey, H. L., Ellis, J. J., Tallent, W. H., and Wolff, I. A., 1969, Mycotoxins as a possible cause of fescue toxicity, J. Agric. Food Chem. 17:437-442. Zimmerman, J. L., Carlton, W. W., and Tuite, J., 1979, Mycotoxicosis produced by cultural products of an isolate of Aspergillus ochraceus. 1. Clinical observations and pathology, Vet. Pathol. 16:583-592.

RECOMMENDATIONS CONCERNING THE CHRONIC PROBLEM OF MISIDENTIFICATION OF MYCOTOXIGENIC FUNGI ASSOCIATED WITH FOODS AND FEEDS Jens C. Frisvad, Kristian F. Nielsen and Robert A. Samson*

1.

INTRODUCTION

Since the aflatoxins were first reported in 1961 from Aspergillus flavus, mycotoxins have often been named after the fungus which was first found to produce them. A long list of connections between fungal species and mycotoxins and antibiotics has been reported, but unfortunately many of the identifications, and hence the connection between mycotoxin name and the source of the toxin, are incorrect (Frisvad, 1989). The most famous example of such incorrect connections was Alexander Fleming’s identification of the original penicillin producer as Penicillium rubrum. Fortunately, in this example, the substance was named after the genus Penicillium, rather than the species, as K. B. Raper re-identified the strain as P. notatum, which was subsequently determined to be a synonym of P. chrysogenum (Pitt, 1979b). Later, penicillin was found in other strains of P. chrysogenum (Raper and Thom, 1949). The early aflatoxin literature is plagued with wrong reports of aflatoxin production by Penicillium puberulum (Hodges et al., 1964), * J. C. Frisvad and K. F. Nielsen: BioCentrum-DTU, Building 221, Technical University of Denmark, 2800 Lyngby, Denmark; R. A. Samson: Centraalbureau voor Schimmelcultures, PO Box 85167, 3508 AD, Utrecht, Netherlands. Correspondence to: [email protected]

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Aspergillus ostianus (Scott et al., 1967), Rhizopus sp. (Kulik and Holaday, 1966), the bacterium Streptomyces (Mishra and Murthy, 1968) and several other taxa. The most famous of these reports was the paper of El-Hag and Morse (1976). They reported that Aspergillus oryzae, the domesticated species used in the manufacture of soy sauce and other Oriental fermented foods, produced aflatoxin. However, the culture of A. oryzae they used was quickly shown to be contaminated by an aflatoxin producing A. parasiticus (Fennell, 1976). Immediate correction of this error did not prevent Adebajo et al. (1992), El-Kady et al. (1994), Atalla et al. (2003) or Drusch and Ragab (2003) reporting that A. oryzae produces aflatoxin. Often, publications reporting mycotoxin production are reviewed by people who have little or no understanding of mycological taxonomy. For example, “P. patulinum” and “P. clavatus” are mentioned in Drusch and Ragab (2003). In Bhatnagar et al. (2002), “P. niger” is mentioned as producing ochratoxin A. Each of these names is an incorrect combination of genus and species. Bhatnagar et al. (2002) give P. viridicatum as producing ochratoxin A in a table, while using P. verruculosum as the species name in the text, confusing it with P. verrucosum, the correct name for the producer of this toxin. Such mistakes could have been avoided. This paper provides a set of recommendations to be followed to ensure correct reports of connections between mycotoxin production and fungal species.

2.

EXAMPLES OF INCORRECT CITATIONS OF SOME FUNGI PRODUCING WELL KNOWN MYCOTOXINS

2.1.

Aflatoxin

The known producers of aflatoxin are given in a separate paper in these Proceedings (Frisvad et al., 2006). The list of other species that have been (incorrectly) reported to produce aflatoxins includes Aspergillus flavo-fuscus, A. glaucus, A. niger, A. oryzae, A. ostianus, A. sulphureus, A. tamarii, A. terreus, A. terricola, A. wentii, Emericella nidulans (as A. nidulans), Emer. rugulosa (as A. rugulosus), Eurotium chevalieri, Eur. repens, Eur. rubrum, Mucor mucedo, Penicillium citrinum, P. citromyces, P. digitatum, P. frequentans, P. expansum, P. glaucum, P. puberulum, P. variabile, Rhizopus sp. and the bacterium Streptomyces sp. None of

Recommendations Concerning the Chronic Problem

35

these species produce aflatoxins, and many of these names are not accepted as valid species in any case.

2.2.

Sterigmatocystin

Fungi known to produce sterigmatocystin include Aspergillus versicolor, Emericella nidulans, several other Emericella species and some Chaetomium species. Although sterigmatocystin is a precursor of aflatoxins (Frisvad, 1989), only Aspergillus ochraceoroseus (Frisvad et al., 1999; Klich et al., 2000), and some Emericella species accumulate both sterigmatocystin and aflatoxin (Frisvad et al., 2004a; Frisvad and Samson, 2004a). Species in Aspergillus section Flavi, which includes the major aflatoxin producers, efficiently convert sterigmatocystin into 3-methoxysterigmatocystin and then into aflatoxins (Frisvad et al., 1999). Many Aspergillus species have been reported to produce sterigmatocystin, incorrectly except for those cited above. Sterigmatocystin production by Penicillium species has not been reported, apart from an obscure reference to Penicillium luteum (Dean, 1963). However, Wilson et al. (2002) claimed that P. camemberti, P. commune and P. griseofulvum produce sterigmatocystin. Perhaps they mistook sterigmatocystin for cyclopiazonic acid. Three Eurotium species have been claimed to produce sterigmatocystin (Schroeder and Kelton, 1975), but this was based only on unconfirmed TLC assays. Unfortunately the strains used were not placed in a culture collection.

2.3.

Ochratoxin A

Ochratoxin A is produced by four main species, Aspergillus carbonarius, A. ochraceus, Petromyces alliaceus, Penicillium verrucosum, and a few other related species as detailed elsewhere (Frisvad and Samson, 2004b; Samson and Frisvad, 2004; Frisvad et al., 2006). A very large number of species have been claimed to produce ochratoxin A, but not all will be detailed here. However, some of the names frequently cited in reviews will be mentioned. Of the Penicillia, P. viridicatum was the name cited for many years as the major ochratoxin A producer, but it was shown that P. verrucosum was the correct name for this fungus, the only species that produces ochratoxin A in cereals in Europe (Frisvad and Filtenborg,1983; Frisvad, 1985; Pitt. 1987). The closely related P. nordicum, which occurs on dried meat in Europe, was mentioned as producing ochratoxin A by Frisvad and Filtenborg (1983) and Land and Hult (1987), but not accepted as a separate species until the publication of Larsen et al. (2001).

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P. verrucosum has been correctly cited as the main Penicillium species producing ochratoxin A for a number of years now, but in a series of recent reviews and papers P. viridicatum and P. verruculosum (no doubt mistaken for P. verrucosum) have been mentioned again (Mantle and McHugh, 1993; Bhatnagar et al., 2002; Czerwiecki et al., 2002a, b). In the latter two papers P. chrysogenum, P. cyclopium, P. griseofulvum, P. solitum, Aspergillus flavus, A. versicolor and Eurotium glaucum were listed as ochratoxin A producers. The strain of P. solitum reported by Mantle and McHugh (1993) to produce ochratoxin A were assigned more recently to P. polonicum, but neither species produces ochratoxin A (Lund and Frisvad, 1994; 2003). These isolates were contaminated by P. verrucosum. The reports by Czerwiecki et al. (2002 a, b) are more problematic in that the fungi have been discarded, so it will never be possible to check the results. The following species were listed as ochratoxin A producers by Varga et al. (2001): Aspergillus auricomus, A. fumigatus, A. glaucus, A. melleus, A. ostianus, A. petrakii, A. repens, A. sydowii, A. terreus, A. ustus, A. versicolor, A. wentii, Penicillium aurantiogriseum, P. canescens, P. chrysogenum, P. commune, P. corylophilum, P. cyaneum, P. expansum, P. fuscum, P. hirayamae, P. implicatum, P. janczewskii, P. melinii, P. miczynskii, P. montanense, P. purpurescens, P. purpurogenum, P. raistrickii, P. sclerotiorum, P. spinulosum,, P. simplicissimum, P. variabile and P. verruculosum. None of these species produces ochratoxin A, and it seems clear that the authors have uncritically accepted lists from earlier reviews. In the recent Handbook of Fungal Secondary Metabolites (Cole and Schweikert, 2003a, b; Cole et al., 2003), only two of the species cited as producing ochratoxin A are correct: A. ochraceus and A. sulphureus. The others mentioned are not.

2.4.

Citrinin

Citrinin is produced by a number of species in Penicillium and Aspergillus, notably P. citrinum, P. expansum, P. verrucosum, A. carneus, A. niveus and an Aspergillus species resembling A. terreus (Frisvad, 1989; Frisvad et al., 2004b), but not by Aspergillus oryzae or P. camemberti, as claimed by Bennett and Klich (2003). Critical checking of the original reports clearly did not occur. Many other species have been claimed to produce citrinin, including A. ochraceus (Mantle and McHugh, 1993), A. wentii (Abu-Seidah, 2002) and Eurotium pseudoglaucum (El-Kady et al., 1994), but either fungus or mycotoxin may have been misidentified in these cases.

Recommendations Concerning the Chronic Problem

2.5.

37

Patulin

A number of species in different genera, notably Penicillium, Aspergillus and Byssochlamys, produce patulin. Among the most efficient producers of patulin are Aspergillus clavatus, A. giganteus, A. terreus, Byssochlamys nivea, P. carneum, P. dipodomyicola, Penicillium expansum, P. griseofulvum, P. marinum, P. paneum and several dung associated Penicillia (Frisvad, 1989; Frisvad et al., 2004b). It is not, however, produced by species in all of the 42 genera listed by Steiman et al. (1989) and Okele et al. (1993). These papers include erroneous statements that Alternaria alternata, Fusarium culmorum, Mucor hiemalis, Trichothecium roseum and many others produce patulin. The production of patulin by Alternaria alternata was later reported by Laidou et al. (2001), and mentioned in a review by Drusch and Ragab (2003). However patulin was not found in hundreds of analyses of Alternaria extracts (Montemurro and Visconti, 1992), or in extracts from more than 200 Alternaria cultures tested by us at the Technical University of Denmark (B. Andersen, personal communication).

2.6.

Penitrem A

Many species have been claimed to produce penitrem A, but most have been misidentifications of Penicillium crustosum (Pitt, 1979; Frisvad, 1989). Names given to isolates that were in fact P. crustosum include P. cyclopium, P. verrucosum var. cyclopium, P. verrucosum var. melanochlorum, P. viridicatum, P. commune, P. lanosum, P. lanosocoeruleum, P. granulatum, P. griseum, P. martensii, P. palitans and P. piceum (Frisvad, 1989). Other species which do produce penitrem A include P. carneum, P. melanoconidium, P. tulipae, P. janczewskii, P. glandicola and P. clavigerum (Frisvad et al., 2004b). Only the first three of these species are likely to occur in foods.

2.7.

Cyclopiazonic Acid

Cyclopiazonic acid is produced by Aspergillus flavus, A. oryzae, A. tamarii, A. pseudotamarii, Penicillium camemberti, P. commune, P. dipodomyicola, P. griseofulvum and P. palitans (Goto et al., 1996; Huang et al., 1994; Pitt et al., 1986; Polonelli et al., 1987; Frisvad et al., 2004b). Cyclopiazonic acid was originally isolated from and named after P. cyclopium CSIR 1082, but this fungus was reidentified as P. griseofulvum (Hermansen et al., 1984; Frisvad, 1989). Despite this, most reviews still cite P. cyclopium or P. aurantiogriseum [of which

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Pitt (1979) considered P. cyclopium to be a synonym] as producers (Scott, 1994; Bhatnagar, 2002; Bennett and Klich, 2003). Scott (1994) drew an incorrect conclusion “α-cyclopiazonic acid is a metabolite of several Penicillium and Aspergillus species and is of Canadian interest from two viewpoints. First, one of the important producers (P. aurantiogriseum, formerly P. cyclopium, Pitt et al., 1986), commonly occurs in stored Canadian grains...”

Although P. aurantiogriseum no doubt occurs in cereal grains, it is not a producer of cyclopiazonic acid. Another example of an error being cited repeatedly is the claimed production of cyclopiazonic acid by Aspergillus versicolor (Ohmomo et al., 1973; cited by Bhatnagar et al., 2002) even though Domsch et al. (1980) and Frisvad (1989) had stated that the isolate described by Ohmomo et al. (1973) was correctly identified as A. oryzae, a wellknown producer of cyclopiazonic acid (Orth, 1977). Penicillium hirsutum, P. viridicatum, P. chrysogenum, P. nalgiovense, Aspergillus nidulans and A. wentii have also wrongly been claimed to produce cyclopiazonic acid (Cole et al., 2003; Abu-Seidah, 2003).

2.8.

Xanthomegnin, Viomellein and Vioxanthin

Xanthomegnin, viomellein and vioxanthin are nephrotoxins produced by all members of Aspergillus section Circumdati (Frisvad and Samson, 2000), Penicillium cyclopium, P. freii, P. melanoconidium, P. tricolor and P. viridicatum (Lund and Frisvad, 1994), and by P. janthinellum and some other genera and species which do not occur in foods. Some of these Penicillium species occur in cereals, so these toxins have been found occurring naturally (Scudamore et al., 1986). These toxins are not produced, however, by P. crustosum as reported by Hald et al. (1983), by P. oxalicum as reported by Lee and Skau (1981) or by A. nidulans, A. flavus, A. oryzae or A. terreus as reported by Abu-Seidah (2003).

2.9.

Penicillic Acid

Penicillic acid is associated with Penicillium series Viridicata and Aspergillus section Circumdati (Lund and Frisvad, 1994; Frisvad and Samson, 2000; Frisvad et al., 2004). Production reported by P. roqueforti (Moubasher et al., 1978; Olivigni and Bullman, 1978) is now considered to be due to the similar species P. carneum (Boysen et al., 1996).

Recommendations Concerning the Chronic Problem

2.10.

39

Rubratoxins

Rubratoxins are hepatoxic mycotoxins known to be produced only by the rare species Penicillium crateriforme (Frisvad, 1989). Rubratoxins are not produced by P. rubrum, P. purpurogenum or Aspergillus ochraceus as reported by Moss et al. (1968), Natori et al. (1970) and Abu-Seibah (2003).

2.11.

Trichothecenes

Trichothecenes are especially troublesome as it is only after the introduction of capillary gas chromatography coupled to mass spectrometry (MS) and more recently the introduction of liquid chromatography combined with atmospheric ionization MS that reliable methods have been available for these mycotoxins. Because immunochemical methods have been improved in recent years they also can now be considered valid. However results from TLC and HPLC based methods are dubious, unless combined with immunoaffinity cleanup, as many authors have neglected very time consuming but crucial clean-up steps. Trichothecene have been reported to be produced by several Fusarium species as detailed elsewhere in these proceedings (Frisvad et al., 2006). Marasas et al. (1984) showed that Fusarium nivale, which gave nivalenol its name, does not produce trichothecenes. However, under its newer, correct name, Microdochium nivale was still incorrectly cited as a trichothecene producer in a recent review (Bhatnagar et al., 2002). It has even been claimed recently that Aspergillus species (A. oryzae, A. terreus, A. parasiticus and A. versicolor) produce nivalenol, deoxynivalenol and T-2 toxin (Atilla et al., 2003). A. parasiticus was claimed to produce very high amounts of deoxynivalenol and T-2 toxin after growth on wheat held at 80% relative humidity for 1-2 months. These data are totally implausible. Possibly the wheat was already contaminated with trichothecenes before use, but the high levels indicate that there may have been false positives as well.

3.

RECOMMENDATIONS

To avoid incorrect reporting of fungal species producing particular mycotoxins, we recommend the following rules when working with mycotoxin producing fungi and the reporting of the results:

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3.1. ●

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Jens C. Frisvad et al.

Ensure correct identification and purity of fungal isolates

Fungal isolates from the particular substrate should be checked with the literature on the mycobiota of foods, e.g. Filtenborg et al. (1996), Pitt and Hocking (1997) or Samson et al. (2004), which correlate particular fungal species with particular food types or substrates. Unusual findings especially should be carefully checked. For example, Aspergillus oryzae is the domesticated form of A. flavus and A. sojae is the domesticated form of A. parasiticus, and these fungi are not expected to be isolated other than from production plants used for making Oriental foods or enzymes. Use typical cultures as reference for comparison, both for identification and mycotoxin production. Frisvad et al. (2000), lists typical cultures for each species of common foodborne Penicillium subgenus Penicillium species. Some effective mycotoxin producing cultures are listed in Table 1. Check the purity of cultures, as contaminated cultures are a very common problem. Check for contaminants by growing cultures on standard media such as CYA (Pitt and Hocking, 1997). Especially when fungi are grown on cereals or liquid cultures it is very difficult to assess if the culture is pure, and it necessary to streak them out on agar substrates where it is much easier to see if the culture is pure.

Table 1. Reference cultures for the production of the more common Aspergillus and Penicillium mycotoxins Mycotoxin Producing species and reference culture Aflatoxins B1 and B2 Aspergillus parasiticus CBSa 100926 Aspergillus flavus CBS 573.65 Aflatoxins G1and G2 Aspergillus parasiticus CBS 100926 Sterigmatocystin Aspergillus versicolor CBS 563.90 Ochratoxin A Petromvces alliaceus CBS 110.26 Penicillium verrucosum CBS 223.71 Patulin Aspergillus clavatus CBS 104.45 Penicillium griseofulvum CBS 295.97 Cyclopiazonic acid Penicillium griseofulvum CBS 295.97 Roquefortine C Penicillium griseofulvum CBS 295.97 Citrinin Penicillium citrinum CBS 252.55 Penicillium verrucosum CBS 223.71 Penicillic acid Penicillium cyclopium CBS 144.45 Penitrem A Penicillium crustosum CBS 181.89 Verrucosidin Penicillium polonicum CBS 101479 Xanthomegnin Penicillium cyclopium CBS 144.45 Rubratoxin B Penicillium crateriforme CBS 113161 a CBS = Culture collection of the Centraalbureau voor Schimmelcultures, Utrecht, Netherlands

Recommendations Concerning the Chronic Problem ●

If unusual producers are found, check them carefully for purity and correct identity using the references cited above. A specialist taxonomist may be consulted.

3.2. ●

●

Ensure optimal conditions for mycotoxin production are used

Use several media and growth conditions to ensure that the fungus can actually produce the mycotoxins. Four good media for mycotoxin production are listed in Table 2.

3.5.

●

Ensure substrate is sterile and not already contaminated with mycotoxins

If natural substrates, such as cereals, are used for mycotoxin production, they should be sterilised before use (e.g. by autoclaving or by gamma-irradiation). Control assays should be carried out for all mycotoxins being studied on the material intended for use as the substrate. This will ensure false positives are not reported. Also check for interfering peaks. Natural substrates such as grains may contain interfering compounds, and the chemical composition of these matrices may change during fungal growth. In such matrices, highly selective cleanup procedures should be used and combined with highly selective analytical methods.

3.4. ●

Ensure that cultures are deposited in a recognised culture collection

Deposit all interesting strains producing mycotoxins in international culture collections, and cite the culture collection numbers in any publications regarding the strains. This procedure should be mandatory for all microbial, biochemical and chemical journals.

3.3. ●

41

Ensure appropriate analytical and confirmatory procedures for mycotoxin extraction and identification

Sample preparation methods are important and should be validated. Sample preparation is specific for the food matrix it is designed for. Use only validated analytical methods.

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Table 2. Efficient media for mycotoxin production Czapek Yeast Autolysate agar (CYA) Yeast Extract Sucrose agar (YES) (Pitt, 1979; Pitt and Hocking, 1997) (Frisvad and Filtenborg, 1983) NaNO3 3g Yeast extract (Difco) 20 g K2HPO4 1g Sucrose 150 g KCl 0.5 g MgSO4 • 7H2O 0.5 g MgSO4 • 7H2O 0.5 g ZnSO4 • 7H2O 0.01 g FeSO4 • 7H2O 0.01 g CuSO4 • 5H2O 0.005 g ZnSO4 • 7H2O 0.01 g Agar 20 g CuSO4 • 5H2O 0.005 g Distilled water l litre Yeast extract (Difco) 5g Sucrose 30 g Agar 20 g Distilled water l litre Rice powder Corn steep agar Mercks Malt Extract (MME) agar (RC) (Bullerman, 1974) (El-Banna and Leistner, 1988) Rice powder 50 g Malt extract 30 g Corn steep liquid 40 g Soy peptone 3g ZnSO4 • 7H2O 0.01 g ZnSO4 • 7H2O 0.01 g CuSO4 • 5H2O 0.005 g CuSO4 • 5H2O 0.005 g Agar 20 g Agar 20 g Distilled water l litre Distilled water l litre pH 5.6

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●

●

Use efficient extraction techniques, for example, fumonisins are very polar and penitrem A is very apolar. Extractions should be validated by recovery experiments. Use authenticated standards of the mycotoxins for comparison, ideally as internal and external standards. More than one separation technique should be use, combined with selective detection principles. Single UV, refractive index, evaporative light scattering, or flame ionisation detection are non-specific. Fluorescence and full UV spectra are specific to some compounds, while mass spectrometry and especially tandem mass spectrometry is very selective for most compounds when monitoring several ions. Generally four identification points should give a very specific detection, e.g. obtained by LC-MS/MS monitoring two fragmentation reactions. Use more than one discretionary test to secure correct identification of the mycotoxin, Combined these with derivatization or alternative clean-up procedures when finding unexpected results.

Recommendations Concerning the Chronic Problem

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REFERENCES

Abu-Seidah, A. A., 2003, Secondary metabolites as co-markers in the taxonomy of Aspergilli, Acta Microbiol. Pol. 52:15-23. Adebajo, L. O., 1992, Spoilage moulds and aflatoxins from poultry feeds, Nahrung 36:523-529. Atalla, M. M., Hassanein, N. M., El-Beih, A. A., and Youssef, Y. A.-G., 2003, Mycotoxin production in wheat grains by different Aspergilli in relation to different humidities and storage periods, Nahrung 47:6-10. Bennett, J. W., and Klich, M. A., 2003, Mycotoxins, Clin. Microbiol. Rev. 16:497-516. Bhatnagar, D., Yu, J., and Ehrlich, K. C., 2002, Toxins of filamentous fungi, in: Fungal Allergy and Pathogenicity, M. J. Breitenbach, R. Crameri, and S. B. Lehrer, eds, Chem. Immunol. 81:167-206. Boysen, M., Skouboe, P., Frisvad, J.C., and Rossen, L., 1996, Reclassification of the Penicillium roqueforti group into three species on the basis of molecular genetic and biochemical profiles, Microbiology 142: 541-549. Bullerman, L. B., 1974, A screening medium and method to detect several mycotoxins in mold cultures, J. Milk Food Technol. 37:1-3. Cole, R. J., and Schweikert, M. A. 2003. Handbook of Secondary Fungal Metabolites. Vol. 1, Academic Press, New York. Cole, R. J., and Schweikert, M. A. 2003. Handbook of Secondary Fungal Metabolites. Vol. 2, Academic Press, New York. Cole, R. J., Jarvis, B. B., and Schweikert, M. A., 2003, Handbook of Secondary Fungal Metabolites. Vol. 3, Academic Press, New York. Czerwiecki, L., Czajkowska, D., and Witkowska-Gwiazdowska, A., 2002a, On ochratoxin A and fungal flora in Polish cereals from conventional and ecological farms. Part 1: Occurrence of ochratoxin A and fungi in cereals in 1997, Food Addit. Contam. 19:470-477. Czerwiecki, L., Czajkowska, D., and Witkowska-Gwiazdowska, A., 2002b, On ochratoxin A and fungal flora in Polish cereals from conventional and ecological farms. Part 2: Occurrence of ochratoxin A and fungi in cereals in 1998, Food Addit. Contam. 19:1051-1057. Dean, F. M., 1963, Naturally Occurring Oxygen Compounds, Butterworth, London, p. 526. Drusch, S., and Ragab, W., 2003, Mycotoxins in fruits, fruit juices, and dried fruits, J. Food Prot. 66:1514-1527. Domsch, K. H., Gams, W., and Anderson, T.-H., 1980, Compendium of Soil Fungi, Academic Press, London. El-Banna, A. A., and Leistner, L., 1988, Production of penitrem A by Penicillium crustosum isolated from foodstuffs, Int. J. Food Microbiol. 7:9-17. El-Hag, N., and Morse, R. E., 1976, Aflatoxin production by a variant of Aspergillus oryzae (NRRL 1988) on cowpeas (Vigna sinensis), Science 192:1345-1346. El-Kady, I., El-Maraghy, S., and Zihri, A.-N., 1994, Mycotoxin producing potential of some isolates of Aspergillus flavus and Eurotium groups from meat products, Microbiol. Res. 149:297-307. Fennell, D. I., 1976, Aspergillus oryzae (NRRL strain 1988): a clarification, Science 194:1188. Filtenborg, O., Frisvad, J. C., and Thrane, U., 1996, Moulds in food spoilage, Int. J. Food Microbiol. 33:85-102.

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Frisvad, J. C. 1985. Classification of asymmetric Penicillia using expressions of differentiation, in Advances in Penicillium and Aspergillus systematics, R. A. Samson and J. I. Pitt, eds, Plenum Press, New York, pp. 327-333. Frisvad, J. C. 1989. The connection between the Penicillia and Aspergilli and mycotoxins with special emphasis on misidentified isolates, Arch. Environ. Contam. Toxicol. 18:452-467. Frisvad, J. C., and Filtenborg, O., 1983, Classification of terverticillate Penicillia based on profiles of mycotoxins and other secondary metabolites, Appl. Environ. Microbiol. 46:1301-1310. Frisvad, J. C., and Samson, R. A., 2000, Neopetromyces gen. nov. and an overview of teleomorphs of Aspergillus subgenus Circumdati, Stud. Mycol. 45:201-207. Frisvad, J.C., and Samson, R.A., 2004a, Emericella venezuelensis, a new species with stellate ascospores producing sterigmatocystin and aflatoxin B1, System. Appl. Microbiol. 27:672-680. Frisvad, J.C., and Samson, R.A., 2004b, New ochratoxin producing species of Aspergillus section Circumdati, Stud. Mycol. 50:23-43. Frisvad, J. C., Filtenborg, O., Lund, F., and Samson, R. A., 2000, The homogeneous species and series in subgenus Penicillium are related to mammal nutrition and excretion, in: Integration of Modern Taxonomic Methods for Penicillium and Aspergillus Classification. R. A. Samson and J. I. Pitt, eds, Harwood Academic Publishers, Amsterdam, pp. 265-283. Frisvad, J. C., Houbraken, J., and Samson, R. A., 1999, Aspergillus species and aflatoxin production: a reappraisal, in: Food Microbiology and Food Safety into the Next Millennium, A. C. J. Tuijtelaars, R. A. Samson, F. M. Rombouts and S. Notermans, eds, Foundation Food Micro ‘99, Zeist, Netherlands. pp. 125-126. Frisvad, J. C., Samson, R. A., and Smedsgaard, J., 2004a, Emericella astellata, a new producer of aflatoxin B1, B2, and sterigmatocystin, Lett. Appl. Microbiol. 38:440-445. Frisvad, J. C., Smedsgaard, J., Larsen, T. O., and Samson, R. A., 2004b, Mycotoxins, drugs and other extrolites produced by species in Penicillium subgenus Penicillium, Stud. Mycol. 49:201-241. Frisvad, J. C., Thrane, U., Samson, R. A. and pitt, J. I., 2006, Important mycotoxins, and fungi which produce them, in: advances in Food Mycology, A. D. Hocking, J. I. Pitt, R. A. Samson and U. Thrane, eds, Springer, New York, pp. 3–25. Goto, T., Wicklow, D. T., and Ito, Y, 1996, Aflatoxin and cyclopiazonic acid production by a sclerotium-producing Aspergillus tamarii strain, Appl. Environ. Microbiol. 62:4036-4038. Hald, B., Christensen, D. H., and Krogh, P., 1983, Natural occurrence of the mycotoxin viomellein in barley and the associate quinone-producing Penicillia, Appl. Environ. Microbiol. 42:446-449. Hermansen, K., Frisvad, J. C., Emborg, C., and Hansen, J., 1984, Cyclopiazonic acid production by submerged cultures of Penicillium and Aspergillus strains, FEMS Microbiol. Lett. 21:253-261. Hodges, F. A., Zust, J. R., Smith, H. R., Nelson, A. A., Armbrecht, B. H., and Campbell, A. D., 1964, Mycotoxins: aflatoxin produced by Penicillium puberulum, Science 145:1439. Huang, X., Dorner, J. W., and Chu, F. S., 1994, Production of aflatoxin and cyclopiazonic acid by various Aspergilli: an ELISA approach, Mycotox. Res. 10:101-106. Ito, Y., Peterson, S. W., Wicklow, D. T., and Goto, T., 2001, Aspergillus pseudotamarii, a new aflatoxin producing species in Aspergillus section Flavi, Mycol. Res. 105:233-239.

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Klich, M., Mullaney, E. J., Daly, C. B., and Cary, J. W., 2000, Molecular and physiological aspects of aflatoxin and sterigmatocystin biosynthesis by Aspergillus tamarii and A. ochraceoroseus, Appl. Microbiol. Biotechnol. 53:605-609. Kulik, M. M. and Holaday, C. E., 1966, Aflatoxin: a metabolic product of several fungi, Mycopath. Mycol. Appl. 30:137-140. Laidou, I. A., Thanassoulopoulos, C. C., and Liakopoulou-Kyriakidis, M., 2001, Diffusion of patulin in the flesh of pears inoculated with four post-harvest pathogens, J. Phytopathol. 149:457-461. Land, C. J., and Hult, K., 1987, Mycotoxin production by some wood-associated Penicillium spp., Lett. Appl. Microbiol. 4:41-44. Larsen, T. O., Svendsen, A., and Smedsgaard, J., 2001, Biochemical characterization of ochratoxin A-producing strains of the genus Penicillium, Appl. Environ. Microbiol. 67:3630-3635. Lee, L. S., and Skau, D. B., 1981, Thin layer chromatographic analysis of mycotoxins: a review of recent literature, J. Liquid Chromatogr. 4, Suppl. 1:43-62. Lund, F., and Frisvad, J. C., 1994, Chemotaxonomy of Penicillium aurantiogriseum and related species, Mycol. Res. 98:481-492. Lund, F., and Frisvad, J. C., 2003, Penicillium verrucosum in cereals indicates production of ochratoxin A. J. Appl. Microbiol. 95:1117-1123. Mantle, P. G., and McHugh, K. M., 1993, Nephrotoxic fungi in foods from nephropathy households in Bulgaria, Mycol. Res. 97:205-212. Marasas, W. F. O., Nelson, P. E., and Toussoun, T. A., 1984, Toxigenic Fusarium Species. Identity and Mycotoxicologv, Pennsylvania State University Press, University Park, Pennsylvania. Mishra, S. K., and Murthy, H. S. R., 1968, An extra fungal source of aflatoxins, Curr. Sci. (Mysore) 37:406. Montemurro, N., and Visconti, A., 1992, Alternaria metabolites -chemical and biological data, in: Alternaria. Biology, Plant Diseases and Metabolites, J. Chelkowski and A. Visconti, eds, Elsevier, Amsterdam, pp. 449-557. Moss, M. O., Robinson, F. V. and Wood, A. B., 1968, Rubratoxin B, a toxic metabolite of Penicillium rubrum, Chemy Ind. 1968:587-588. Moubasher, A. H., Abdel-Kader, M. I. A., and El-Kady, I. A., 1978, Toxigenic fungi isolated from Roquefort cheese, Mycopathologia 66:187-190. Natori, S., Sakaki, S., Kurata, M., Udagawa, S., Ichinoe, M., Saito, M., Umeda, M., and Ohtsubo, K., 1970, Production of rubratoxin B by Penicillium purpurogenum, Appl. Microbiol. 19:613-617. Ohmomo, S., Sugita, M., and Abe, M., 1973, Isolation of cyclopiazonic acid, cyclopiazonic acid imine and bissecodehydrocyclopiazonic acid from the cultures of Aspergillus versicolor (Vuill.) Tiraboschi, J. Agric. Chem. Soc. Japan 47:57-93. Okeke, B., Seigle-Murandi, F., Steiman, R., Benoit-Guyod, J.-L., and Kaouadjii, M., 1993, Identification of mycotoxin-producing fungal strains: a step in the isolation of compounds active against rice fungal diseases, J. Agric. Food Chem. 41:1731-1735. Olivigni, F. J., and Bullerman, L. B., 1978, Production of penicillic acid and patulin by an atypical Penicillium roqueforti isolate, Appl. Microbiol. 35:435-438. Orth, R., 1977, Mycotoxins of Aspergillus oryzae strains for use in the food industry as starters and enzyme producing molds, Ann. Nutr. Aliment. 31:617-624. Pitt, J. I., 1979a, Penicillium crustosum and P. simplicissimum, the correct names for two common species producing tremorgenic mycotoxins, Mycologia 71:1166-1177.

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Pitt, J. I., 1979b, The Genus Penicillium and its Teleomorphic States Eupenicillium and Talaromyces, Academic Press, London. Pitt, J. I., 1987., Penicillium viridicatum, P. verrucosum, and the production of ochratoxin A, Appl. Environ. Microbiol. 53:266-269. Pitt, J. I., and Hocking, A. D., 1997, Fungi and Food Spoilage. 2nd edition, Blackie Academic and Professional, London. Pitt, J. I., Cruickshank, R. H., and Leistner, L., 1986, Penicillium commune, P. camembertii, the origin of white cheese moulds, and the production of cyclopiazonic acid, Food Microbiol. 3:363-371. Polonelli, L., Morace, G., Rosa, R., Castagnola, M., and Frisvad, J. C., 1987, Antigenic characterization of Penicillium camemberti and related common cheese contaminants, Appl. Environ. Microbiol. 53:872-878. Raper, K. B. and Thom, C., 1949, A Manual of the Penicillia, Williams and Wilkins, Baltimore. Samson, R. A., and Frisvad, J. C., 2004, New ochratoxin or sclerotium producing species in Aspergillus section Nigri, Stud. Mycol. 50:45-61. Samson, R. A., Hoekstra, E. S., and Frisvad, J. C., eds, 2004, Introduction to Foodand Airborne Fungi, 7th edition, Centraalbureau voor Schimmelcultures, Utrecht, Netherlands, 389 pp. Schroeder, H. W., and Kelton, W. H., 1975, Production of sterigmatocystin by some species of the genus Aspergillus and its toxicity to chicken embryos, Appl. Microbiol. 30:589-591. Scott, P. M., 1994, Penicillium and Aspergillus toxins, in: Mycotoxins in Grain. Compounds other than Aflatoxin. J. D. Miller and H. L. Trenholm, H. L., eds, Eagan Press, St. Paul, Minnesota, pp. 261-285. Scott, P. M., van Walbeek, W., and Forgacs, J., 1967, Formation of aflatoxins by Aspergillus ostianus Wehmer, Appl. Microbiol. 15:945. Scudamore, K. A., Atkin, P., and Buckle, A. E., 1986, Natural occurrence of the naphthoquinone mycotoxins, xanthomegnin, viomellein and vioxanthin in cereals and animal foodstuffs, J. Stored Prod. Res. 22:81-84. Steiman R., Seigle-Murandi, F., Sage, L., and Krivobok S., 1989, Production of patulin by micromycetes, Mycopathologia 105:129-133. Varga, J., Rigó, K., Réren, J., and Mesterházy, Á., 2001, Recent advances in ochratoxin research. I. Production, detection and occurrence of ochratoxins, Cereal Res. Commun. 29:85-92. Wilson, D. M., Mutabanhema, W., and Jurjevic, Z., 2002, Biology and ecology of mycotoxigenic Aspergillus species as related to economy and health concerns, in: Mycotoxins and Food Safety, J. W. DeVries, M. W. Trucksess and L. S. Jackson, eds, Kluwer Academic Publishers, Dordrech, Netherlands. pp. 3-17.

Section 2. Media and method development in food mycology Comparison of hyphal length, ergosterol, mycelium dry weight and colony diameter for quantifying growth of fungi from foods Marta H. Taniwaki, John I. Pitt, Ailsa D. Hocking and Graham H. Fleet Evaluation of molecular methods for the analysis of yeasts in foods and beverages Ai Lin Beh, Graham H. Fleet, C. Prakitchaiwattana and Gillian M. Heard Standardization of methods for detecting heat resistant fungi Jos Houbraken and Robert A. Samson

COMPARISON OF HYPHAL LENGTH, ERGOSTEROL, MYCELIUM DRY WEIGHT, AND COLONY DIAMETER FOR QUANTIFYING GROWTH OF FUNGI FROM FOODS M. H. Taniwaki, J. I. Pitt, A. D. Hocking and G. H. Fleet*

1.

INTRODUCTION

Fungi are significant environmental microorganisms, as they are responsible for spoilage of foods, production of mycotoxins and in some cases desirable bioconversions. It is important therefore to have reliable, convenient methods for measuring fungal growth. However, the growth of fungi is not easy to quantify because, unlike bacteria and yeasts, fungi do not grow as single cells, but as hyphal filaments that cannot be quantified by the usual enumeration techniques. Fungal hyphae can penetrate solid substrates, such as foods, making their extraction difficult. In addition, fungi differentiate to produce spores, resulting in large increases in viable counts often with little relationship to biomass (Pitt, 1984). A number of methods have been developed for quantifying fungal growth and their principles and applications comprehensively reviewed (Matcham et al.,1984; Hartog and Notermans, 1988; Williams, 1989; Newell, 1992; Samson et al., 1992; de Ruiter et al., 1993; Pitt and Hocking, 1997). The most frequently used method is * M. H. Taniwaki, Instituto de Tecnologia de Alimentos, Campinas-Sp, Brazil; J. I. Pitt, A. D. Hocking, Food Science Australia, PO Box 52, North Ryde, NSW 2113, Australia; G. H. Fleet, Food Science and Technology, University of New South Wales, Sydney, NSW 2052, Australia. Correspondence to: [email protected]

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the counting of viable propagules, i.e. colony forming units (CFU), a technique derived from food bacteriology. However, this method suffers from serious drawbacks. Viable counts usually reflect spore numbers rather than biomass (Pitt, 1984). When fungal growth consists predominantly of hyphae, i.e. in young colonies or inside food particles, viable counts will be low, but when sporulation occurs, counts often increase rapidly without any great increase in biomass. Some fungal genera, e.g. Alternaria and Fusarium, produce low numbers of spores in relation to hyphal growth, whereas others, e.g. Penicillium, produce very high numbers of spores. Consequently, viable counts are a poor indicator of the extent of fungal growth and appear to correlate poorly with other measures such as ergosterol (Saxena et al., 2001). A second commonly used method is measurement of colony diameter (Brancato and Golding, 1953). When measured over several time intervals, colony diameters can be translated into growth rates, which are frequently linear over quite long periods (Pitt and Hocking, 1977) and have been widely used in water activity studies (e.g. Pitt and Hocking, 1977; Pitt and Miscamble, 1995) and to model growth (Gibson et al., 1994). However colony diameter as a measure of fungal biomass takes no account of colony density (Wells and Uota, 1970). Estimation of mycelium dry weight is a third commonly used method to assess fungal growth or biomass. This is the method of choice for growth in liquid systems, such as fermentors, however, mycelium dry weight measurements lack sensitivity and are destructive (Deploey and Fergus, 1975). A fourth approach, measurement of hyphal length, was used by Schnürer (1993) to estimate the biomass of three fungi grown in pure culture. This technique has the advantage of actually measuring growth, but is particularly laborious. None of these methods can be used to estimate fungal biomass in foods. Chemical assays have also been used to measure fungal growth. The two substances commonly assayed are chitin and ergosterol. The chitin assay is well documented (Ride and Drysdale, 1972), but has major disadvantages: it lacks sensitivity, is time consuming, and is subject to interference from insect fragments (Pitt and Hocking, 1997). Ergosterol is the dominant sterol in most fungi (Weete, 1974), and is not found to any significant extent in plants, animals or bacteria (Schwardorf and Muller, 1989). Thus, its measurement in environmental samples can be taken as an index of the presence of fungi (Seitz et al., 1977, 1979; Nylund and Wallander, 1992; Miller and

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Young, 1997). The advantages of this method are high sensitivity, specificity and relatively short analysis time (Seitz et al., 1977; Schwardorf and Muller, 1989). However, the ergosterol assay has never been validated against more traditional methods, an essential step before it can be accepted as a reliable method for quantifying fungal growth in foods. In addition, the influences of such factors as medium composition, water activity and age of colony on the ergosterol content of mycelium have not been evaluated adequately. One study has attempted to compare ergosterol content with mycelial dry weight over a range of species: for nine aquatic fungi, only three showed correlations between these parameters (Bermingham et al., 1995). Studies comparing ergosterol content with mould viable counts have reported mixed results in grains (Schnürer and Jonsson, 1992) and pure cultures (Saxena et al., 2001). This paper reports a comparison of the ergosterol, colony diameter, dry weight and hyphal length methods for quantifying the growth of several fungal species significant in foods. Studies were carried out in pure culture under a range of conditions.

2.

MATERIALS AND METHODS

2.1.

Fungi

Single isolates of nine food spoilage fungi, representing examples of heat resistant, xerophilic and toxigenic species commonly found in foods, were obtained from the FRR culture collection at Food Science Australia North Ryde, NSW, Australia (Table 1). These species were Aspergillus flavus, Byssochlamys fulva, Byssochlamys nivea, Eurotium chevalieri, Fusarium oxysporum, Mucor plumbeus, Penicillium commune, Penicillium roqueforti and Xeromyces bisporus.

2.2.

Media

The following media were used: Czapek Yeast Extract Agar (CYA), Malt Extract Agar (MEA) and Potato Dextrose Agar (PDA) representative of high water activity (aw) media, (all of ca 0.997 aw); and Czapek Yeast Extract 20% Sucrose Agar (CY20S), 0.98 aw and Malt Extract Yeast Extract 50% Glucose Agar (MY50G), 0.89 aw, as reduced aw media. PDA was from Oxoid Ltd, Basingstoke, UK, and the formulae for the others are given by Pitt and Hocking (1997).

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Table 1. Origins of cultures useda Species Strain number Aspergillus flavus FRR 2757 Byssochlamys fulva FRR 3792 Byssochlamys nivea FRR 4421 Eurotium chevalieri FRR 547 Fusarium oxysporum FRR 3414 Mucor plumbeus FRR 2412 Penicillium commune FRR 3932 Penicillium roqueforti FRR 2162 Xeromyces bisporus FRR 2351 a FRR denotes the culture collection Australia

2.3.

Source Peanut, Queensland, Australia, 1984 Strawberry puree, NSW, Australia, 1990 Strawberry, Brazil, 1993 Animal feed, Queensland, Australia, 1970 Orange juice, NSW, Australia, 1987 Apple juice, NSW, Australia, 1981 Cheddar cheese, NSW, Australia, 1991 Cheddar cheese, USA, 1978 Dates, NSW, 1981 of Food Science Australia, North Ryde, NSW,

Cultivation

Inocula were prepared from 5 to 7 day cultures grown on CYA, except for B. nivea, E. chevalieri and X. bisporus which were grown on MEA for 7 to 10 days, CY20S for 10 to 15 days and MY50G for 15 to 20 days, respectively. Cultures for growth estimates and assays were grown in 90 mm plastic Petri dishes, inoculated at a single central point. Each fungus was grown on several plates of each medium. Plates were incubated upright at 25°C.

2.4.

Growth Measurement

Fungal growth was measured by the methods described below throughout the growth period, but at intervals which varied widely with species and medium. Measurements and assays were carried out in duplicate. 2.4.1.

Colony Diameters

Colonies were measured from the reverse side in millimetres with a ruler. Only well formed, circular colonies were chosen for measurement. 2.4.2.

Mycelium Dry Weight

A colony and surrounding agar were cut from a Petri dish, transferred to a beaker containing distilled water (100 ml), then heated in a

Comparison of Hyphal Length, Ergosterol, Mycelium Dry Weight

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steamer for 30 min to melt the agar. The mycelium, which remained intact, was rinsed once in distilled water and then transferred to a dried, weighed filter paper which was placed in an aluminium dish and dried in an oven at 80°C for 18 h. After cooling to room temperature in a desiccator, the filter papers and mycelium were weighed and the dry weight calculated by difference. The method was based on those of Paster et al. (1983) and Zill et al. (1988). 2.4.3.

Hyphal Length

Hyphal lengths were estimated by direct microscopy using a haemocytometer and a modification of the method of Schnürer (1993). Colonies and associated agar were cut into pieces and homogenized with distilled water (3-100 ml, depending on the size of the colony), for about 30 s using an Ultra-Turrax homogeniser (Ystral GmbH, Dottingen, Germany). The suspension was then treated in a sonicator (Branson Sonic Power Company, Danbury, CT) at 100 watts for about 20 s to break up hyphal clumps. After dilution in distilled water, drops (0.5 ml) were placed in a haemocytometer and hyphal fragments counted. Hyphal lengths were measured using the intersection technique (Olson, 1950) at a magnification of 400 X. Colonies from two plates were measured separately and for each plate 10 fields were counted. Results were calculated from the means of the two plates. 2.4.4.

Ergosterol Assay

A colony was excised from a Petri dish culture, transferred to a beaker of distilled water (100 ml) containing Tween 80 (0.05%) and steamed for 30 min to melt the agar. The intact mycelium was collected, rinsed with water and transferred to a round bottomed flask. Ergosterol was extracted from the mycelium by refluxing with 95% ethanol: water (100 ml, 50: 50 v/v) and potassium hydroxide (5 g) for 30 min (Zill et al., 1988). This crude extract was partitioned three times with n-hexane in a separating funnel. The combined hexane extracts were concentrated under vacuum to near dryness. The residue was redissolved in n-hexane (2 ml), then filtered through a polypropylene membrane, 0.45 µm pore size, 13 mm diameter (Activon, Sydney, NSW). The filtrate was dried under N2 and redissolved in n-hexane for ergosterol quantification. Ergosterol was assayed by high pressure liquid chromatography (HPLC) using a Millipore Waters system fitted with a LiChrosorb SI

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60 column (Gold Pak, Activon). The column was eluted with nhexane: isopropanol (97: 3, v/v) at 1 ml/min and ergosterol was detected by absorption at 280 nm about 8-10 min after injection of the sample. Ergosterol was quantified by reference to an ergosterol standard calibration curve, prepared from a standard solution (2 mg/ml, Sigma Chemicals, St. Louis, MO). For five of the fungal

Figure 1. HPLC traces of ergosterol from six fungi: (a) Penicillium commune, (b) Penicillium roqueforti, (c) Byssochlamys nivea, (d) Fusarium oxysporum, (e) Aspergillus flavus, (f) Eurotium chevalieri. The ergosterol peak is indicated by an arrow.

Comparison of Hyphal Length, Ergosterol, Mycelium Dry Weight

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species, well separated single peaks for ergosterol were obtained in HPLC traces of mycelial extracts (Figure 1). However, in extracts from E. chevalieri and X. bisporus the peak eluted close to interfering substances. In these cases a second filtration of the extract or addition of ergosterol standard was necessary to conclusively identify the ergosterol peaks.

3.

RESULTS

3.1.

Validation of ergosterol assay

A linear relationship was observed between peak heights measured on HPLC chromatograms and ergosterol concentration. The lower limit of detection was 0.01 µg of ergosterol. The coefficient of variation for ergosterol peaks detected by HPLC in extracts ranged between 1 and 14% provided that the ergosterol content was greater than 20 µg. Variation was greatest when the ergosterol content was less than 50 µg and least for high amounts (e.g. 500 µg) (Table 2). Ergosterol recoveries from spiked samples of F. oxysporum and B. fulva mycelium were 80-100% using the described method. For most species, the ergosterol peak in the HPLC traces was clear and well separated from other peaks (Figure 1a-d). However, the ergosterol peaks for Aspergillus flavus (Figure 1e) and more especially for Eurotium chevalieri (Figure 1f) and Xeromyces bisporus (not shown) were close to peaks likely to be other sterols. Levels of ergosterol observed in these species were lower than expected.

Table 2. Reproducibility of ergosterol analysis (n=3) in colonies of Fusarium oxysporum and Byssochlamys fulva grown on Czapek yeast extract agar for various incubation periods Incubation Coefficient of Species time (d) Ergosterol (µg) variation (%) Fusarium oxysporum 2 5.5, 7.5,11.1 34.9 5 620.5, 654.8, 696.2 5.8 6 884.2, 767.4, 835.0 7.0 Byssochlamys fulva 2 0.76, 0.42, 0.76 28.4 4 23.5, 31.1, 27.7 13.9 7 694.7, 682.0, 686.2 0.9

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M. H. Taniwaki et al.

Growth of Fungi as Assessed by Colony Diameters

Most of the fungi grew on all of the media used, though with varying vigour, reflecting their water relations (Table 3). Best growth of most species occurred on CY20S, 0.98 aw except for B. fulva and F. oxysporum which grew faster at 25°C on CYA, 0.997 aw. Along with M. plumbeus, these species produced little or no growth on MY50G at 0.89 aw, an aw near their lower limit for growth (Pitt and Hocking, 1997). A. flavus and P. commune grew strongly at all aw tested. Growth of E. chevalieri, a xerophilic species, was slow on CYA, and faster on CY20S and MY50G. X. bisporus, an extreme xerophile, grew only on MY50G.

3.3.

Influence of Colony Age on Growth Parameters

The influence of colony age on some of the data obtained by the four methods used for measuring fungal growth is shown in Table 4, for four representative species. Ratios were calculated for colony diameter over hyphal length, mycelium dry weight and ergosterol content over hyphal length, and ergosterol over mycelium dry weight. The ratio of colony diameter over hyphal length showed a general downward trend, indicating a greater rate of hyphal extension than colony diameter increase as colonies aged. Colony diameters are therefore not a good measure of fungal biomass production in aging colonies. The other ratios were reasonably constant, indicating a general correspondence between mycelial dry weight, ergosterol content and hyphal length.

Table 3. Colony diameters of fungi grown on media of various water activities at 25°C Species Colony diameter (mm) at 7 days on CYA 0.997 aw CY20S 0.98 aw MY50G 0.89 aw Aspergillus flavus 68 84 16 Byssochlamys fulva 78 58 0 Byssochlamys nivea 43 -a Eurotium chevalieri 19 50 36 Fusarium oxysporum 90 78 6 Mucor plumbeus 63 85 4 Penicillium commune 32 44 18 Penicillium roqueforti 52 Xeromyces bisporus 0 0 12 a not tested

Comparison of Hyphal Length, Ergosterol, Mycelium Dry Weight

Table 4. Growth of four species of fungi on Czapek yeast extract agar as measured by four techniques, and ratios derived from those measurementsa Hyphal Colony ErgoMycelium length diam sterol Ratio Ratio dry Ratio Ratio Species Time (d) wt (mg) (m ×1000) MDW/HL (mm) (µg) CD/HL E/HL E/MDW 2.04 8.1 Mucor 3 51 11.2 25 5.5 16.5 0.68 2.62 11.8 6 plumbeus 73 62.0 27.9 23.7 30.9 2.00 4.34 8.3 16 83 81.2 19.2 18.7 36.0 2.26 Fusarium 4 10.45 3.9 44 128.6 4.2 12.3 40.4 3.18 6 oxysporum 32.38 4.1 70 444.3 2.1 13.7 133.8 3.32 9 83.32 3.7 86 598.3 1.0 7.1 308.6 1.94 2.51 5.9 Byssochlamys 5 33 77.5 13.1 30.9 14.7 5.27 8 fulva 19.83 6.0 71 379.6 3.6 19.1 119.6 3.17 9 30.75 6.0 86 977.7 2.8 31.8 185.4 5.27 3.23 6.1 Penicillium 4 29 43.6 9.0 13.5 19.6 2.22 8.36 12.3 7 roqueforti 57 168.6 6.8 20.2 103.4 1.63 14 19.22 12.4 86 280.5 4.5 14.6 238.4 1.18 a Ratio CD/HL, ratio of colony diameter (mm) / hyphal length (m ×1000); ratio MDW/HL, ratio of mycelial dry weight (mg) / hyphal length (m ×1000), ratio E/HL, ergosterol (µg) / hyphal length (m ×1000).

57

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M. H. Taniwaki et al.

Estimates of Fungal Growth by Mycelial Dry Weight and Ergosterol Compared with Hyphal Length

To provide a common reference point, colonies of diameter 83-86 mm, i.e. virtually full plate growth from a single inoculum point, were selected where possible. To take account of type of medium, two data sets were developed, for colonies on CYA and PDA. Species other than Penicillium commune, E. chevalieri and X. bisporus produced full plate colonies on both media, although after varied incubation periods. P. commune reached 86 mm on PDA after 21 days, but a maximum of only 39 mm on CYA, after 17 days. E. chevalieri colonies reached a maximum of only 46 mm diameter on PDA, after 16 days, and were smaller on CYA. For comparisons with other species on CYA, therefore, E. chevalieri colonies on CY20S were used. As expected, X. bisporus did not grow on either CYA or CY20S, so data from MY50G were used, where this species reached a maximum of 64 mm after 42 days incubation. The overall results from analyses of hyphal length, mycelium dry weight and ergosterol under these conditions are given in Table 5.

3.5.

Hyphal Length

Hyphal lengths, estimated for colonies of similar diameters as set out in Table 5, varied widely between species. On CYA, hyphal length varied almost 20 fold between F. oxysporum (83,000 m) and B. nivea (4200 m). Results were more uniform on PDA, with less than 6 fold variation among the species. F. oxysporum produced only 23,000 m of hyphae on PDA. B. nivea produced the greatest hyphal length on PDA (28,000 m) but the lowest on CYA (4200 m). Despite profuse growth, M. plumbeus produced less than 6000 m of hyphae under any of the varied conditions used. Mycelial dry weights were also lower for M. plumbeus than for many of the other species studied.

3.6.

Ergosterol Content

Ergosterol content per colony varied widely between genus, between medium and even within genus, reflecting differences in growth density and membrane composition. When grown on CYA, B. fulva produced the highest ergosterol content and M. plumbeus the

Ergosterol (µg)

Ratio E/HL

Ratio E/MDW

46.4 31.7 15.5 7.17 18.7 57.4 14.6 21.4

1.00 5.28 2.74 1.93 2.25 2.75 1.18 2.44

Medium: CYA Aspergillus flavus Byssochlamys fulva B. nivea Fusarium oxysporum Mucor plumbeus Penicillium commune P. roqueforti Average

86 (11) 86 (9) 86 (17) 86 (9) 83 (13) 39 (17) 86 (14)

6.42 30.8 4.17 83.3 4.34 7.67 19.2

297 185 23.5 309 36 160 238

46.3 6.01 5.63 3.71 8.29 20.9 12.4 14.7

298 977 64.5 598 81.2 440 281

86 (11) 86 (7) 86 (9) 46 (16) 86 (7) 86 (4) 86 (2 1) 86 (11)

4.78 22.2 28.4 8.6 23.3 5.2 7.02 7.48

186 156 180 86 171 109 108 222

38.9 7.02 6.33 10.0 7.33 21.0 15.4 29.7 17.0

84.3 1463 183 669 1884 259 818 509

17.6 65.9 6.4 7.8 80.9 49.8 116.5 68 51.6

0.45 9.37 1.02 7.78 11.0 2.36 7.57 2.29 5.23

86 (14)

6.59

354

53.7

25.1

3.8

0.07

Medium: PDA Aspergillus flavus Byssochlamys fulva B. nivea Eurotium chevalieri Fusarium oxysporum Mucor plumbeus Penicillium commune P. roqueforti Average

Medium: CY20S Eurotium chevalieri

Medium: MY50G

59

42.7 Xeromyces bisporus 7.22 64 (42) 28.9 4 10.7 1.47 Overall average 36.4 17.6 4.1 a Ratio CD/HL, ratio of colony diameter (mm)/ hyphal length (m ×1000); ratio MDW/HL, ratio of mycelial dry weight (mg)/ hyphal length (m ×1000), ratio E/HL, ergosterol (µg) /hyphal length (m × 1000).

Comparison of Hyphal Length, Ergosterol, Mycelium Dry Weight

Table 5. Comparison of measurements of mature growth of various fungi on CYAa Colony diameter Hyphal Mycelium mm (incubation Ratio dry length Species weight (mg) time, d) (m ×1000) MDW/HL

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lowest; about a 12 fold difference. On PDA, F. oxysporum produced the highest ergosterol content, with A. flavus the lowest, about a 22 fold difference. For some species, medium composition greatly affected ergosterol content. This was most evident for E. chevalieri, which produced 25 times as much ergosterol on PDA (670 µg/colony) from colonies less than 50 mm in diameter, than from 86 mm diameter colonies on CY20S (25 µg), despite comparable hyphal lengths on the two media. In contrast, ergosterol production by A. flavus was much higher on CYA than on PDA, again with comparable hyphal lengths.

3.7.

Mycelium Dry Weights

Mycelium dry weights of 83-86 mm diameter colonies varied between species. On CYA, A. flavus and F. oxysporum produced colonies with a high mycelium dry weight (approximately 300 mg/plate). On CY20S, E. chevalieri colonies were equally heavy. The weights of M. plumbeus colonies, however were only 12% of these values (Table 5). For the vigorously growing Aspergillus, Penicillium and Fusarium species, mycelium dry weights on PDA were lower than values obtained on CYA. However, mycelium weights for M. plumbeus and B. nivea were much higher on PDA than on CYA. In the case of B. nivea, this difference was more than 8 fold.

3.8.

Relationship Between Hyphal Length and Mycelium Dry Weight

The relationship between hyphal length and mycelium dry weight over time of growth was reasonably constant within species (Table 4) except for very small colonies of Penicillium roqueforti, and varied only four fold between the four species shown in Table 4. When all nine species were compared, on more than one medium, much greater variability was seen. This appears to be due mostly to variation in hyphal length measurements, which is not so precise as the other techniques used here. Increases in hyphal length sometimes occurred with little increase in mycelial dry weight, e.g. for M. plumbeus when grown on CYA and E. chevalieri grown on PDA. These observations were reproducible (data not shown). As discussed by Schnürer (1993), vacuole formation and autolysis of cell contents may occur in aging cultures which would lead to a reduction in weight per unit length.

Comparison of Hyphal Length, Ergosterol, Mycelium Dry Weight

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On the other hand, some species sporulated heavily in age, e.g. P. roqueforti grown on PDA, A. flavus on CYA, and E. chevalieri on CY20S. Here large increases in mycelial dry weight were accompanied by little or no hyphal growth. The Penicillium, Aspergillus, and Eurotium species showed ratios above 12 mg/1000 m, while for the Mucor, Fusarium and Byssochlamys species ratios were 8 mg/1000 m or below. On PDA, ratios ranged from 6.3 mg/1000 m (B. nivea) to 38.9 mg/1000 m (A. flavus).

3.9.

Relationship Between Hyphal Length and Ergosterol Content

The ratios of ergosterol production (µg) to hyphal length (m ×1000) for colonies of various ages were found to be more variable within species than those for mycelial dry weight over hyphal length (Table 4). No pattern with age of cultures was apparent. On PDA, values varied from 6.4 (B. nivea) to 116.5 (P. commune), an 18 fold difference (Table 5). Low ergosterol production on PDA by A. flavus and on CY20S by E. chevalieri (see above) was reflected in very low ratios of ergosterol to hyphal length.

3.10.

Relationship Between Ergosterol Content and Mycelial Dry Weight

Reasonable agreement was seen between the ratios of ergosterol (µg) to mycelium dry weight (mg) both for cultures of different age in each species and between species (Table 4). If the very low value for small colonies of M. plumbeus (0.68) is omitted, ratios varied between 1.18 and 5.27, less than 5 fold. When the effect of medium is considered with the full range of species (Table 5), ratios were again reasonably constant for colonies grown on CYA and PDA. With omission again of a very low figure (0.45 for A. flavus on PDA), values varied from 1.0 to 11.0. The average for all species was 4.1 (Table 5). Most species produced higher amounts of ergosterol and mycelium dry weight on PDA than on CYA except for A. flavus, which produced higher ergosterol and mycelium dry weight on CYA than on PDA. However, the very low ratio (0.07) observed from growth of E. chevalieri on CY20S is anomalous. No doubt this is due to the very low level of ergosterol produced on this medium by this species. The HPLC spectrum showed several peaks eluting close to ergosterol, so other sterols may have been present.

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DISCUSSION

This study attempted to validate ergosterol measurements as an index of fungal growth for important food spoilage fungi, chosen because of the great differences in their growth patterns, using hyphal length and mycelium dry weight as standards. Colony diameter, hyphal length, mycelium dry weight and ergosterol content all gave useful information about the growth of the species examined. However, each technique exhibited advantages and limitations. Colony diameter is a sensitive technique, in that a colony as small as 2 mm is easily measured. It was not possible to determine accurately dry weight, hyphal length or ergosterol concentration on such small amounts of material. However, colony diameter did not show a consistent correlation with the other parameters, especially as colony diameters became larger and more mature. In particular, sporulation caused a reduced correlation between colony diameters and the other parameters. If colony diameters are measured from relatively early growth, e.g. using the Petrislide technique of Pitt and Hocking (1977), then correlations could be higher. Values for hyphal length obtained from single colonies on standard Petri dishes by Schnürer (1993) ranged from 10,000 m for Rhizopus stolonifer to 54,000 m for Fusarium culmorum. In this study, comparable fungi produced more hyphae: 43,000 m for Mucor plumbeus and 83,000 m for Fusarium oxysporum. These differences may be due to the lower nutritional value of the medium used by Schnürer (1993). In this study, the average ratio of mycelium dry weight over hyphal length was 17.6 mg dry weight/1000 m of mycelium. Schnürer (1993) found values of 4.2 to 6.7 mg/1000 m, calculated from the mycelium and hyphal volume, respectively. Values obtained by us for Fusarium oxysporum, Byssochlamys spp. grown on CYA and PDA, and Mucor plumbeus when grown on CYA were comparable with those of Schnürer (1993). Much larger differences were seen with the rapidly growing and/or highly sporulating Aspergillus and Penicillium species studied here. Given the differences in fungi studied and media used, the data of Schnürer (1993) and our data are comparable. Working with aquatic fungi, ratios of ergosterol to mycelium dry weight of 2.3 to 11.5 were given by Gessner and Chauvet (1993), similar to those of Schnürer (1993). Reports have suggested that ergosterol content increases as colonies age (Nout et al., 1987; Torres et al., 1992). However, as discussed by Schnürer (1993), vacuole formation and autolysis of cell contents may occur in aging cultures which would lead to a reduction in weight per

Comparison of Hyphal Length, Ergosterol, Mycelium Dry Weight

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unit of length and, consequently, to an increased ergosterol to dry weight ratio. Differences in degree and type of sporulation by the species studied here also clearly play a major part in the variations in the ratios of mycelial dry weight to hyphal length and ergosterol content to hyphal length observed here. For example, Aspergillus flavus and the Penicillium species produce relatively little vegetative mycelium relative to conidiophores and conidia, increasing mycelial dry weight and probably decreasing ergosterol in relation to hyphal length. Mycelium dry weight is often considered to be a basic measure of fungal growth but fundamental questions remain unanswered. In this study, mycelium was separated from agar medium using a heat treatment. Cochrane (1958) criticised the separation of agar medium from fungal biomass using hot water because the water may extract soluble fungal components, resulting in a loss of dry weight. However, it is also important to note that dry weights measured without prior extraction are greatly affected by the variation in internal solutes caused by different medium formulations, especially in media of reduced aw (Hocking and Norton, 1983). An alternative approach is to scrape or peel the fungal growth from the medium surface. This may lead to incomplete removal and underestimation of dry weight. The technique of Hocking (1986) where fungi were grown on dialysis membrane on the surface of agar media enables ready separation of fungus from medium, and is a notable improvement. However, in the current study, where some species sporulated profusely, a wet extraction technique was considered preferable for safety reasons. In this study, extracted mycelium dry weight showed a reasonably good correlation with hyphal length (Tables 4, 5), indicating the value of this parameter as a measure of fungal growth in media. However, the measurement of mycelium dry weight is not readily applicable to the estimation of growth of fungi in foods. Schnürer (1993) noted differences in growth patterns between different fungal species. For a nonsporulating Fusarium culmorum, good agreement was found between hyphal length, colony counts and ergosterol content. For Penicillium rugulosum and Rhizopus stolonifer, changes in ergosterol level were related more closely to changes in hyphal length rather than to production of spores or colony counts. In the present study, hyphal length correlated rather poorly with ergosterol content (Table 5). Schnürer and Jonsson (1992) found reasonable correlation between ergosterol and mould viable counts in Swedish grains under certain conditions, but Saxena et al. (2001) found that

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viable counts and ergosterol did not correlate well for pure cultures of Aspergillus ochraceus and Penicillium verrucosum. The accuracy of the hyphal length technique is affected by factors including variation in hyphal width for different species, degree of sporulation, formation of reproductive structures (e.g. cleistothecia), fragmentation during homogenization, and clumping of hyphae. The intersection technique of Olsen (1950) is probably only statistically sound when large numbers of microscopic fields are counted. Despite the laboriousness of the procedure used here for hyphal length estimations (ten fields from two colonies), this rigour is lacking. All of these factors lead to variation in estimates of hyphae length. Ergosterol content was a sensitive indication of fungal biomass. As little as 0.01 µg of ergosterol could be detected from mycelium in a colony of 4 mm diameter. However, the amount of ergosterol found in the fungi varied with the growth medium, species and culture incubation time. This variation was reflected in the data shown in Tables 4 and 5. The ratio of ergosterol over mycelium dry weight ranged from 0.07 µg/mg for E. chevalieri on CY20S to 11.0 µg/mg for F. oxysporum on PDA, a 150 fold variation. Even when these two extreme values are omitted, variation of about 20 fold remained (i.e. 0.45 µg/mg for A. flavus to 9.37 µg/mg for B. fulva, both on PDA). Weete (1974) noted that in general sterol levels in fungi varied with medium composition and culture conditions. Four to 10 fold variations have been reported in the ergosterol content of the same fungus under different growth conditions (Newell et al., 1987; Nout et al., 1987). Increased nutritional complexity of the medium, the presence of free fatty acid precursors of the ergosterol biosynthetic pathway and increased availability of oxygen all gave mycelium with increased ergosterol contents. The measurement of ergosterol alone does not give the absolute amount of fungus present. For this, it is necessary to convert ergosterol values into biomass in terms of mycelium dry weight. After studying 14 aquatic hyphomycetes, Gessner and Chauvet (1993) gave the range of ratios of ergosterol to mycelium dry weight as 2.3 -11.5 µg/g, figures similar to those derived in this work (0.45 -11.0 µg/g). This ratio varies between fungal species and with growth condition, limiting the direct use of ergosterol as a means of calculating mycelium dry weight. A further explanation for variation in the ergosterol content of fungi is that sterols other than ergosterol can be produced by some species. For example, ergosterol and 22-dihydroergosterol have been reported as the predominant sterol in A. flavus (Vacheron and Michel,

Comparison of Hyphal Length, Ergosterol, Mycelium Dry Weight

65

1968; Weete, 1973). Other sterols identified as products of deuteromycetous fungi include cerevisterol, ergosterol peroxide, lanosterol, 24-methylenelophenol and 14-dehydroergosterol (Weete, 1973). In this study, A. flavus produced a low level of ergosterol despite its high production of biomass, and showed extra peaks in its extract. Similarly, colonies of E. chevalieri were found to contain only low amounts of ergosterol, despite high amounts of biomass. Additional peaks seen in the HPLC profile of E. chevalieri extracts also indicate that it may produce sterols other than ergosterol. The ergosterol content of X. bisporus mycelium was also low, and several additional peaks were observed in the HPLC trace from its extract. Further studies are needed to determine if the additional peaks found in the HPLC profiles are sterols. The low level of ergosterol produced by E. chevalieri has important consequences. Eurotium species are very common in stored grains, in which ergosterol has been used to estimate fungal growth. The overall average ratio of ergosterol to mycelium dry weight was 4.1, about 60 times higher than that obtained for E. chevalieri on CY20S. Hypothetically, a sample of grain infected by E. chevalieri, estimated to contain a particular ergosterol content, could contain 150 times as much fungal biomass as one infected by Fusarium oxysporum, which gave an ergosterol to mycelium dry weight ratio of 11 µg/mg in this study. Estimation of ergosterol content, colony diameter, mycelium dry weight and hyphal length were shown to be good indices for measuring fungal growth, but it is important to keep in mind the limitations of each techniques. The most reliable information about fungal growth will be obtained by using two or more techniques for quantification.

5.

ACKNOWLEDGMENTS

The authors wish to thank to Mr N. Tobin and Ms S. L. Leong of Food Science Australia, North Ryde, for helpful advice on chemical analyses and hyphal length measurement, respectively, and to Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP) for funding the PhD program for M.H.T.

6.

REFERENCES

Bermingham, S., Maltby, L., and Cooke, R. C., 1995, A critical assessment of the validity of ergosterol as an indicator of fungal biomass, Mycol. Res. 99:479-484.

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Brancato, F. P., and Golding, N. S., 1953, The diameter of the mold colony as a reliable indicator of growth, Mycologia 45:848-864. Cochrane, V. W., 1958, Physiology of Fungi, John Wiley, New York. Deploey, J. J., and Fergus, C. L., 1975, Growth and sporulation of thermophilic fungi and actinomycetes in O2-N2 atmospheres, Mycologia 67:780-797. de Ruiter, G. A., Notermans, S. H. W., and Rombouts, F. M., 1993, New methods in food mycology, Trends Food Sci. Technol. 4:91-97. Gessner, M. O. and Chauvet, E., 1993, Ergosterol-to-biomass conversion factors for aquatic Hyphomycetes, Appl. Environ. Microbiol. 59:502-507. Gibson, A. M., Baranyi, J., Pitt, J. I., Eyles, M. J., and Roberts, T. A., 1994, Predicting fungal growth: the effect of water activity on Aspergillus flavus and related species, Int. J. Food Microbiol. 23:419-431. Hartog, B. J. and Notermans, S., 1988, The detection and quantification of fungi in food, in: Introduction to Food-borne Fungi. 4th edition, R. A. Samson and E. S. van Reenen-Hoestra, eds, Centraalbureau voor Schimmelcultures, Baarn, Netherlands, pp. 222-230. Hocking, A. D., 1986, Some Physiological Responses of Fungi Growing at Reduced Water Activities, PhD thesis, University of New South Wales, Kensington, NSW. Hocking, A. D., and Norton, R. S., 1983, Natural abundance 13C nuclear magnetic resonance studies on the internal solutes of xerophilic fungi, J. Gen. Microbiol. 129:2915-2925. Matcham, S. E., Jordan, B. R., and Wood, D. A., 1984, Methods for assessment of fungal growth on solid substrates, in: Microbiological Methods for Environmental Biotechnology. J. M. Grainger and J. M. Lynch, eds, Academic Press, London, pp. 5-18. Miller, J. D., and Young, J. C., 1997, The use of ergosterol to measure exposure to fungal propagules in indoor air, Am. Ind. Hyg. Assoc. J. 58:39-43. Newell, S. Y., 1992, Estimating fungal biomass and productivity in decomposing litter, in: The Fungal Community, G. C. Carroll and D. T. Wicklow, eds, Marcel Dekker, Inc., New York, pp. 521-561. Newell, S. Y., Miller, J. D., and Fallon, R. D., 1987, Ergosterol content of salt-marsh fungi: effect of growth conditions and mycelial age, Mycologia 79:688-695. Nout, M. J. R., Bonants-van Laarhoven, T. M. G., de Jongh, P., and Koster, P. G., 1987, Ergosterol content of Rhizopus oligosporus NRRL 5905 grown in liquid and solid substrates, Appl. Microbiol. Biotechnol. 26:456-461. Nylund, J. E., and Wallander, H., 1992, Ergosterol analysis as a means of quantifying mycorrhizal biomass, in: Methods in Microbiology, Vol 24, Techniques for the Study of Mycorrhiza. J. R. Norris, D. J. Read, and A. K. Varma, eds, Academic Press, London, pp.77-88. Olson, F. C. W., 1950, Quantitative estimates of filamentous algae, Trans. Am. Microsc. Soc. 69:272-279. Paster, N., Lisker, N., and Chet, I., 1983, Ochratoxin A production by Aspergillus ochraceus Wilhelm grown under controlled atmospheres, Appl. Environ. Microbiol. 45:1136-1139. Pitt, J. I., 1984, The significance of potentially toxigenic fungi in foods, Food Technol. Aust. 36:218-219. Pitt, J. I., and Hocking, A. D., 1977, Influence of solute and hydrogen ion concentration on the water relations of some xerophilic fungi, J. Gen. Microbiol. 101:35-40. Pitt, J. I., and Hocking, A. D., 1997, Fungi and Food Spoilage, 2nd edition, Blackie Academic and Professional, London.

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Pitt, J. I., and Miscamble, B. F., 1995, Water relations of Aspergillus flavus and closely related species, J. Food Prot. 58:86-90. Ride, J. P., and Drysdale, R. B., 1972, A rapid method for the chemical estimation of filamentous fungi in plant tissue, Physiol. Plant Pathol. 2:7-15. Samson, R. A., Hocking, A. D., Pitt J. I., and King, A. D., 1992, Modern Methods in Food Mycology, Elsevier Publishers, Amsterdam. Saxena, J., Munimbazi, C., and Bullerman, L. B., 2001, Relationship of mould count, ergosterol and ochratoxin A production, Int. J. Food Microbiol. 71: 29-34. Schnürer, J., 1993, Comparison of methods for estimating the biomass of three foodborne fungi with different growth patterns, Appl. Environ. Microbiol. 59:552-555. Schnürer, J., and Jonsson, A., 1992, Ergosterol levels and mould colony forming units in Swedish grains of food and feed grade, Acta Agric. Scand. Sect B. 42:240-245. Schwardorf, K., and Muller, H. M., 1989, Determination of ergosterol in cereals, mixed feed components, and mixed feeds by liquid chromatography, J. Assoc. Off. Anal. Chem. 72:457-462. Seitz, L. M., Mohr, H. E., Burroughs, R., and Sauer D. B., 1977, Ergosterol as an indicator of fungal invasion in grain, Cereal Chem. 54:1207-1217. Seitz, L. M., Sauer, D. B., Burroughs, R., Mohr, H. E., and Hubbard J. D., 1979, Ergosterol as a measure of fungal growth, Phytopathology 69:1202-1203. Torres, M., Viladrich, R., Sanchis, V., and Canela, R., 1992, Influence of age on ergosterol content in mycelium of Aspergillus ochraceus, Lett. Appl. Microbiol. 15:20-22. Vacheron, M. J., and Michel, G., 1968, Composition en sterols et en acides gras de deux souches d’Aspergillus flavus, Phytochemistry 7:1645-1651. Weete, J. D., 1973, Sterols of fungi: distribution and biosynthesis, Phytochemistry 12:1843-1864. Weete, J. D., 1974, Distribution of sterols in the fungi. 1. Fungal spores, Lipids 9: 578-581. Wells, J. M., and Uota, M., 1970, Germination and growth of five fungi in low-oxygen and high-carbon dioxide atmospheres, Phytopathologia 60:50-53. Williams, A. P., 1989, Methodological developments in food mycology, J. Appl. Bacteriol. 67: Symp. Suppl. 61S-67S. Zill, G., Engelhardt, G., and Wallnofer, P. R., 1988. Determination of ergosterol as a measure of fungal growth using Si 60 HPLC, Z. Lebensm. Unters. Forsch. 187: 246-249.

EVALUATION OF MOLECULAR METHODS FOR THE ANALYSIS OF YEASTS IN FOODS AND BEVERAGES Ai Lin Beh, Graham H. Fleet, C. Prakitchaiwattana and Gillian M. Heard *

1.

INTRODUCTION

The analysis of yeasts in foods and beverages involves the sequential operations of isolation, enumeration, taxonomic identification to genus and species, and strain differentiation. Although well established cultural methods are available to perform these operations, many molecular methods have now been developed as alternatives. These newer methods offer various advantages, including faster results, increased specificity of analysis, decreased workload, computer processing of data and possibilities for automation. Molecular methods for yeast analysis are now at a stage of development where they can move from the research laboratory into the quality assurance laboratories of the food and beverage industries. However, many practical questions need to be considered for this transition to progress. A diversity of molecular methods with similar analytical objectives are available. Which methods should the food analyst choose and what principles should be used to guide this choice? Food analysts are required to make judgements and decisions about the microbiological quality and safety of consignments of products often worth many millions of dollars in national and international trade. Moreover, * Food Science and Technology, School of Chemical Engineering and Industrial Chemistry, University of New South Wales, Sydney, New South Wales, Australia, 2052. Correspondence to: [email protected]

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these decisions need to conform to the requirements of supplier and customer contracts, and government legislation. Consequently, they have the potential to encounter intense legal scrutiny (Fleet 2001). For these reasons, the food analyst requires basic information about method standardisation, accuracy, reproducibility, precision, specificity and detection sensitivity (Cox and Fleet, 2003). While these criteria have guided the selection and choice of currently accepted cultural methods, they have not been critically applied to the newer molecular techniques. This Chapter has the following goals: (1) to provide an overview of the diversity of molecular methods that are finding routine application to the analysis of yeasts in foods and beverages, (ii) to outline the variables that affect the performance and reliability of these methods, and (iii) to suggest strategies for the international standardisation and validation of these methods. Molecular methods have found most application to the identification of yeast species and to strain differentiation, but there is increasing use of culture-independent methods to detect and monitor yeasts in food and beverage ecosystems.

2.

MOLECULAR METHODS FOR YEAST IDENTIFICATION

The traditional, standard approach to yeast identification has been based on cultural, phenotypic analyses. The yeast isolate is examined for a vast range of morphological, biochemical and physiological properties which are systematically compared with standard descriptions to give a genus and species identity. Generally, it is necessary to conduct approximately 100 individual tests to obtain a reasonably reliable identification (Kurtzman and Fell, 1998; Barnett et al., 2000). Consequently, the entire process is very labour-intensive, lengthy and costly. Although various technical and diagnostic innovations have been developed to facilitate this process, they are not universal in their application and the data generated are not always equivalent (Deak and Beuchat, 1996; Deak 2003; Robert, 2003; Kurtzman et al., 2003). Molecular methods based on DNA analysis are now being used to quickly identify yeasts to genus and species level. The workload is minimal and, usually, reliable data can be obtained within 1-2 days. Several approaches are being used. The most definitive and universal assay determines the sequence of bases in segments of the ribosomal DNA. Other approaches are based on determination of restriction fragment

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length polymorphisms (RFLP) of segments of ribosomal DNA, hybridisation with specific nucleic acid probes, and polymerase chain reaction (PCR) assays with species-specific primers. Aspects of these methods and their application to food and beverage yeasts have been reviewed by Loureiro and Querol (1999), Giudici and Pulvirenti (2002), Loureiro and Malfeito-Ferreira (2003), and van der Vossen et al. (2003).

2.1.

Sequencing of Ribosomal DNA

The discovery that ribosomal RNA is highly conserved throughout nature but has certain segments which are species variable, has lead to the widespread use of ribosomal DNA sequencing in developing microbial phylogeny and taxonomy. As a consequence, ribosomal DNA sequences are known for most microorganisms, including yeasts, and are now routinely used for diagnostic and identification objectives (Valente et al., 1999). The ribosomal DNA repeat unit found in yeasts is schematically shown in Figure 1. It consists of conserved and variable regions which are arranged in tandem repeats of several hundred copies per genome. The conserved sequences are found in genes encoding for small (18S), 5.8S, 5S, and large (25-28S) subunits of ribosomal RNA. Within each cluster, variable spacer regions occur between the subunits, called internal transcribed spacer (ITS) regions, and between gene clusters, called the intergenic spacer regions (IGS) or the non-transcribed spacer region (NTS). All of these regions have some potential for differentiating yeast genera and species, but most focus has been on the 18S, 26S and ITS regions (Valente et al., 1999; Kurtzman, 2003). The D1/D2 domain of the large subunit (26S) ribosomal DNA consists of about 600 nucleotides and has been sequenced for virtually all known yeast species. Databases of these sequences can be accessed through GenBank (http://www.ncbi.nlm.nih.gov/), DataBank of Japan (http://www.ddbj.nig.ac.jp/) or the European Molecular Biology Laboratory (http://www.ebi.ac.uk/embl/). There is sufficient variation in these sequences to allow differentiation of most ascomycetous (Kurtzman and Robnett, 1998, 2003) and basidiomycetous (Fell et al.

ss rDNA 18S

ITS

ITS

5.8SS

ls rDNA 26S

Figure 1. The ribosomal DNA repeat unit

NTS

NTS 5S

SS rDNA 18S

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2000; Scorzetti et al., 2002) yeast species. Sequencing of the D1/D2 domain of the 26S ribosomal DNA is now widely used for the routine identification of yeasts and the construction of phylogenetic taxonomy. Sequence comparisons have also been done for the small subunit, 18S ribosomal DNA but, so far, the databases are not extensive and sequence differences may not be sufficient to allow the discrimination of closely related species (James et al., 1997; Naumov et al., 2000; Daniel and Meyer, 2003). The ribosomal spacer regions (ITS) show higher rates of sequence divergence than the D1/D2 domain of the 26S subunit and have proven useful for species differentiation (James et al., 1996; Naumov et al., 2000; Cadez et al., 2003). For example, the Hanseniaspora uvaurm-guillermondii cluster is poorly resolved, and species of Saccharomyces pastorianus/Saccharomyces bayanus, and Cryptococcus magnus/Filobasidium floriforme/Filobasidium elegans are indistinguishable using D1/D2 sequences. Sequencing of the ITS region can provide the required level of differentiation. Sequencing of mitochondrial and protein encoding genes are also being used to determine phylogenetic relationships among yeasts. These genes include the translation elongation factor 1α, actin-1, RNA polymerase II, pyruvate decarboxylase, beta tubulin gene, small subunit rDNA and cytochrome oxidase II (Daniel et al., 2001; Kurtzman and Robnett 2003; Daniel and Meyer 2003). The basic protocol for sequencing ribosomal DNA segments is: (i) prepare a pure culture of the yeast isolate, (ii) extract and purify the DNA, (iii) perform PCR amplification of the region to be sequenced, (iv) verify the amplified product by gel eletrophoresis, and (v) sequence the product using internal or external primers. Procedures for conducting these operations are well established but are not standardised, and may vary from one laboratory to another. Primer sequences used to amplify the different segments of the rDNA have been tabulated in White et al. (1990), Valente et al. (1999), Sipiczki (2002) and Kurtzman and Robnett (2003). Table 1 lists some key publications on the identification of yeasts by ribosomal DNA sequencing. Some yeasts are not reliably identified by sequencing single gene segments and it is suggested that sequences be obtained for multiple genes or gene segments for more reliable data (Kurtzman, 2003).

2.2.

Restriction Fragment Length Polymorphism (RFLP)

RFLP analysis of the ribosomal DNA segments is emerging as one of the most useful methods for rapidly identifying food and beverage

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Table 1. Application of gene sequencing technology to the identification of species of food and beverage yeasts Region Application References 18S Phylogenetic relationships; Zygosaccharomyces and Torulaspora species James et al. (1994) Brettanomyces, Dekkera, Cai et al. (1996) Debaryomyces, Kluyveromyces species Candida, Pichia, Citeromyces species Suzuki and Nakase (1999) Saccharomyces genus; new species James et al. (1997) S. kunashirensis, S. martiniae Saccharomyces sensu lato group; Mikata et al. (2001) new species S. naganishii, S. humaticus, S. yukushimaensis 18S, ITS Phylogenetic relationships of Naumov et al. (2000) Saccharomyces sensu stricto complex; new species S. cariocanus, S. kudriavzevii, S. mikatae 18S; 834Identification of yeast from Cappa and Cocconcelli (2001) 1415 dairy products D1/D2 Systematics of ascomycetous yeasts Kurtzman and Robnett (1998) of 26S Systematics of basdidiomycetous Fell et al. (2000) yeasts D1/D2 of Systematics of basdidiomycetous Scorzetti et al. (2002) 26S, ITS yeasts D1/D2 of Candida davenportii sp. nov. from Stratford et al. (2002) 26S a wasp in a soft-drink production facility D1/D2 of Tetrapisispora fleetii sp. nov. from Kurtzman et al. (2004) 26S, ITS a food processing plant D1/D2 Identification of yeast species; of 26S in Sicilian sourdough Pulvirenti et al. (2001) in orange juice Arias et al. (2002) in spontaneous wine fermentation van Keulen et al. (2003) from bark of cork oak Villa-Carvajal et al. (2004) from Malbec grape berries Combina et al. (2005) from fermentation of West Jespersen et al. (2005) African cocoa beans contaminant in carbonated orange Pina et al. (2005) juice production chain ITS Phylogenetic relationships of Belloch et al. (2002) Kluyveromyces marxianus group ITS Phylogenetic relationships of James et al. (1996) Zygosaccharomyces and Torulaspora species

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Table 1. Application of gene sequencing technology to the identification of species of food and beverage yeasts—cont’d Region Application References ITS Phylogenetic analysis of the Oda et al. (1997) Saccharomyces species Phylogenetic analysis of the Montrocher et al. (1998) Saccharomyces sensu stricto complex ITS Identification of yeast species; in orange fruit and orange juice Heras-Vazques et al. (2002) from Italian sourdough baked Foschino et al. 2004 products ITS1 Separation of S. cerevisiae strains Naumova et al. 2003 in African sorghum beer IGS Separation of Clavispora opuntiae Lachance et al. 2000 varieties ITS, IGS Intraspecies diversity of Mrakia Diaz and Fell 2000 and Phaffia species Actin Phylogenetic relationships of Daniel et al. 2001 anamorphic Candida and related teleomorphic genera mt COX II Phylogeny of the genus Belloch et al. 2000 Kluyveromyces Multigenes Ascomycete phylogeny Kurtzman and Robnett Ascomycete species separation (2003), Kurtzman (2003), Daniel and Meyer (2003) Multigenes Taxonomic position of the van der Aa Kühle and biotherapeutic agent Jespersen (2003) Saccharomyces boulardii

yeasts. The preferred region for analysis represents the ITS1-5.8SITS2 segment (Figure 1). Using appropriate primers, this segment is specifically amplified by PCR. The PCR product is then cleaved with specific restriction endonucleases, and the resulting fragments are separated by gel electrophoresis. The size (number of base pairs) of the ITS amplicon itself can be useful in discriminating between yeast species. The number (usually 1-4) and size (base pairs) of the fragments as determined by banding patterns on the gel are the main principles used to discriminate between yeast species. Generally, more than one restriction enzyme needs to be used in order to obtain unequivocal discrimination. Some restriction enzymes commonly used are: Cfo I, Hae III, Hinf I, Hpa II, Scr FI, Taq I, Nde II, Dde I, Dra I and Mbo II. Several hundred species of food and beverage yeasts have now been examined by this method and databases of fragment profiles for the different restriction enzymes and yeast species have been established

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(Esteve-Zarzoso et al., 1999; Granchi et al., 1999; Arias et al., 2002; Heras-Vazquez et al., 2003; Dias et al., 2003; Naumova et al., 2003). PCR-RFLP analyses have several advantages that are attractive to quality assurance analysis in the food and beverage industries. Once a pure yeast culture has been obtained, identification to species level can be done in several hours. Essentially, DNA is extracted from the yeast biomass, amplified by specific PCR, amplicons are digested with the restriction nucleases and the products separated by gel electrophoresis. The work load and equipment needs are minimal and data are generally reproducible. The expense and time for sequencing are avoided. Although PCR-RFLP analysis of the ITS1-5.8S-ITS2 region has attracted most study to date, there is increasing interest in the PCRRFLP analysis of other ribosomal segments. These include the 18SITS region, 18S-ITS-5.8S region, the 26S and NTS regions. It is not evident at this stage whether targeting these regions offers any advantage over the ITS1-5.8S-ITS2 region and further studies evaluating the different approaches are required. Table 2 lists some key reports on the application of the PCR-RFLP analysis of ribosomal DNA regions to food and beverage yeasts.

2.3.

Nucleic acid probes and species-specific primers

Nucleic acid probes are short, single-stranded nucleotides (usually 20-100 bases) that are designed to complement a specific sequence in the DNA/RNA of the target organism. They are usually labelled with a marker molecule to enable their detection. Probes are used in hybridization reactions, and are applied in a number of formats (Hill and Jinneman, 2000; Cox and Fleet, 2003). In whole cell hybridization protocols (FISH, CISH), yeasts are directly visualised in situ, and identified with fluorescently-labelled, specific probes that bind to rRNA, located in the ribosomes. In ecological studies, this technique is particularly useful for identifying morphological types, for quantifying target species and monitoring microbial community structure and dynamics, for example, in examining the spatial relationships on surfaces of leaves and in biofilms (Table 3). Probes are also used in dot blot, slot blot and colony blot hybridisation assays of yeast biomass on membranes. Detection is achieved by a labeled DNA probe that hybridises to the DNA/RNA of the immobilised sample (Hill and Jinneman, 2000). Another strategy is to coat the probe onto a solid substrate such as the wells of a microtitre tray, and use a modified ELISA format to detect the target

76 Ai Lin Beh et al.

Table 2. Application of PCR-restriction fragment length polymorphism (RFLP) to the identification of food and beverage yeasts Method: region; primers; restriction enzymes References Applications ITS2; (primers ITS3 and ITS4); AseI, Chen et al. (2000) Medically-important yeast species BanI, EcoRI, HincII, StyI ITS1-5.8S rRNA-ITS2; (primers ITS1 Guillamón et al. (1998), Ganga and Yeast species from wine fermentations and ITS4); CfoI, HaeIII, HinfI Martinez. (2004), Combina et al. (2005) Yeast species from Irish cider fermentations Morrissey et al. (2004) Yeast species associated with orange juice Arias et al. (2002) ITS1-5.8S rRNA-ITS2; (primers ITS1 Esteve-Zarzoso et al. (1999) 132 species from food and beverages and ITS4); CfoI, HaeIII, HinfI, DdeI Granchi et al. (1999), Rodríguez et al. Yeast species from wine fermentations (2004), Clemente-Jimenez et al. (2004) Esteve-Zarzoso et al. (2001) Yeast species during fermentation and ageing of sherry wines Yeast species from orange fruit and orange juice Heraz-Vazquez et al. (2003) 1) ITS1-5.8S rRNA-ITS2; (primers ITS1 Pulvirenti et al. (2001) Yeast species from Sicilian sourdoughs and ITS4); HinfI, RsaI, NdeII, HaeIII 2) NTS 2; (primers r-1234 and r-2156); AluI, BanI ITS1-5.8S rRNA-ITS2; (primers ITS1 Saccharomyces sensu stricto strains from Naumova et al. (2003) African sorghum beer and ITS4); HaeIII, HpaII, ScrFI, TaqI ITS1-5.8S rRNA-ITS2; (primers ITS1 Antunovics et al. (2005) Differentiation of S. bayanus, S. cerevisiae, and ITS4); HaeIII, MaeI S. paradoxus isolates from botrytised grape must ITS1-5.8S rRNA-ITS2; (primers ITS1 Cadez et al. (2003) Species within the genera of Hanseniaspora and Kloeckera and ITS4); HinfI, DdeI, MboII

Baleiras Couto et al. (1996a)

128 species from food, wine, beer and soft drinks

Dlauchy et al. (1999)

Yeast species from Hungarian dairy products

Vasdinyei and Deak (2003)

Discrimination of C. stellata, M. pulcherrima, K. apiculata and S. pombe

Capece et al. (2003)

Differentiation of S. cerevisiae and S. paradoxus isolates from Croatian vineyards Separation of Saccharomyces sensu stricto and Torulaspora species Differentiation of brewery yeasts; S. carlsbergensis, S. pastorianus, S. bayanus, S. cerevisiae, S. brasiliensis, S. exiguus Discrimination of S. cerevisiae, S. carlsbergensis and S. pastorianus Differentiation of S. uvarum and S. cerevisiae from wine Separation of S. bayanus, S. cerevisiae and S. paradoxus wine strains Yeast species from wine fermentations Yeast species from wine fermentations

Redzepovic et al. (2002) Smole Mozina et al. (1997) Barszczewski and Robak (2004)

Molina et al. (1993) Demuyter et al. (2004) Antunovics et al. (2005) van Keulen et al. (2003) Baleiras Couto et al. (2005)

77

26S rRNA; (primers NL1 and NL4); AluI 26S rRNA; (primers NL1 and LRS); MseI, ApaI, HinfI

Discrimination/diversity of S. cerevisiae strains

Evaluation of Molecular Methods for the Analysis of Yeasts in Foods

1) ITS; (primers ITS1 and NL2); MseI, TaqI 2) NTS; (primers JV51ET and JV52ET); MseI, TaqI 18S rRNA-ITS1; (primers NS1 and ITS2); HaeIII, MspI, AluI, RsaI 18S rRNA-ITS1; (primers NS1 and ITS2); HaeIII, MspI 1) 18S rRNA; (primers p108 and M3989); HaeIII, MspI 2) NTS; (primers NTSF and NTSR); HaeIII, MspI 18SrDNA and ITS1; (primers NS1 and ITS2); HaeIII, MspI 18SrDNA and ITS1; (primers NS1 and ITS2); CfoI, HaeIII, HinfI, MspI ITS1-5.8S rRNA-ITS2-part 18SrRNA ; (primers NS3 and ITS4); ScrFI, HaeIII, MspI 3′ ETS and IGS; (primers 5S2 and ETS2); MspI, ScrFI MET2 ; EcoRI, PstI

D. bruxellensis isolates from winery air samples

Connell et al. (2002)

CISH; PNA probes in 18S rRNA and 26S rRNA

S. cerevisiae, Z. bailii, D. bruxellensis colonies on filter membranes

Perry-O’Keefe et al. (2001)

FISH; DNA probes in 18S rRNA

S. cerevisiae, P. anomala, D. bruxellensis and D. hansenii isolates, detection in yoghurts

Kosse et al. (1997)

FISH; DNA probes in 18S rRNA

Detection and quantification of A. pullulans on leaf surfaces

Spear et al. (1999), Andrews et al. (2002)

D. hansenii isolates from cheese, differentiation of hansenii and fabryii varieties.

Corredor et al. (2000)

Candida sp. EJ1 in wine samples B. bruxellensis in wine

Mills et al. (2002) Cocolin et al. (2004)

Detection of marine yeast species; P. guillermondii, R. diobovatum, R. sphaerocarpum, K.thermotolerans-like, C. parapsilosis, C. tropicalis, D. hansenii Detection of C. albicans, C. tropicalis, C. krusei in blood Identification of 18 Candida species

Kiesling et al. (2001)

Dot blot/slot blot hybridization Dot blot hybridization RNA slot blot hybridization PCR-ELISA Probes in D1/D2 region Probes immobilized on plates, PCR with biotinylated primers Probes in ITS2 region Probes labelled with DIG Capture PCR amplicons on strepavidin coated plates

Fujita et al. (1995) Elie et al. (1998)

Ai Lin Beh et al.

CISH; PNA probe in D1/D2 26S rRNA

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Table 3. Application of nucleic acid probes and specific primers for the detection of food and beverage yeasts Probe/primer Format References Application Whole cell hybridization; Fluorescent/chemiluminescent in situ hybridization (FISH/CISH) FISH; PNA probe in D1/D2 26S rRNA Stender et al. (2001), Dias et al. (2003) D. bruxellensis isolates from wine

2) one species-specific, the other universal (ITS region) Nested PCR Multiplex PCR 5 primers; 1 universal, 4 species-specific (ITS region) PCR and RT-PCR Specific primers (D1/D2 26S LSU) RT-PCR; Specific primer pairs (cs 1) gene RT-PCR; Specific primers (ITS and LSU region) PCR, Multiplex PCR; 4 primers; 2 species-specific pairs (YBR033w region) Real time PCR Primer pairs (D1/D2 LSU) Primer pairs (rad4 gene) Specific primer pairs (cs 1) gene Universal primer pairs ITS3 and ITS4 (5.8S and ITS2)

Pathogenic yeasts C. neoformans, T. cutaneum, R. mucilaginosa Detection of several Candida species

Fell (1995)

Identification of Z. bailii, Z. bisporus, Z. rouxii and T. delbrueckii isolates from fruit

Sancho et al. (2000)

D. bruxellensis strains from isolates and sherry Identification of Dekkera isolates; differentiation of B. bruxellensis, B. anomala, B. custersianus, B. naardenensis Detection of B. bruxellensis/B. anomalus from wine samples Detection of viable C. krusei in fruit juice Identification of S. cerevisiae and S. bayanus/pastorianus isolates Detection of S. cerevisiae, S. bayanus and S. pastorianus

Ibeas et al. (1996) Egli and Henick-Kling (2001)

Detection and enumeration of D. bruxellensis in wines Detection and quantification of B. bruxellensis in wines Quantification of C. krusei from fruit juice Differentiation of Z. bailii, Z. rouxii, C. krusei, R. glutinis and S. cerevisiae by difference in Tm

Mannarelli and Kurtzman (1998)

Cocolin et al. (2004) Casey and Dobson (2003) Josepa et al. (2000) Torriani et al. (2004)

Phister and Mills (2003) Delaherche et al. (2004) Casey and Dobson (2004) Casey and Dobson (2004)

Evaluation of Molecular Methods for the Analysis of Yeasts in Foods

Species-specific primers Universal and species-specific primers (V3 region of LSU) Universal and species-specific primers (D1/D2 region of LSU) 1) species-specific primer pairs

79

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DNA in PCR amplicons (Kiesling et al., 2002) (Table 3). Speciesspecific primers are used in PCR assays to generate amplicons. Production of the amplicon means that the particular target species is present in the sample. Qualitative detection of the target amplicon is done by its visualisation in gel electrophoresis. Real time PCR systems are now being applied to yeasts, and allow the simultaneous detection and quantification of the target species, omitting the electrophoresis step (Table 3). Nucleic acid probes and specific primers have gained widespread use in the detection of bacterial species. Their application to the detection of yeast species has not been that extensive and further development is needed. Most probes reported to date have been developed around specific sequences in ribosomal DNA, and it would be worthwhile to identify other species-specific genes that could be targeted for probe development.

2.4.

Differentiation of Strains Within a Species

The distinctive character and appeal of many foods and beverages (eg. bread, beer, wine) produced by fermentation with yeasts are frequently attributable to the contribution and properties of particular strains. Strain typing is also useful to trace the source of yeast contamination in outbreaks of food spoilage. The ability to differentiate strains within a species is, therefore, an important requirement in quality assurance programs. Over the past 20 years, a diversity of molecular methods has been developed and applied to the differentiation of yeast strains, and some of these are sufficiently robust and convenient for routine use (Table 4). Electrophoretic karyotyping of genomic DNA using pulse field gel electrophoresis (PFGE) and RFLP analysis of genomic DNA have been widely applied to “fingerprint” yeast strains with very good success and confidence, but they require significant attention to DNA preparation and extraction, as well as to subsequent electrophoretic analyses (Cardinali and Martini, 1994; Deak, 1995; van der Aa Kühle et al., 2001). Analysis of mitochondrial DNA by RFLP produces fragment profiles that give excellent strain discrimination. Simplified methods for extraction and processing of the mitochondrial DNA have greatly improved the convenience and reliability of this assay, and consequently it has found significant application to the analysis of food and beverage yeasts (Querol et al., 1992; López et al., 2001; see review of Loureiro and MalfietoFerreira, 2003).

AFLP EcoRI-C/Mse-AC MseI-C/PstI-AA, -AC, -AT EcoRI/MseI, nine primer pairs MseI-EcoRI four primer pairs EcoRI-C/MseI-AC EcoRI-MseI four primer pairs

Species and strain differentiation of Saccharomyces and non-Saccharomyces wine yeasts

de Barros Lopes et al. (1999, 2002)

Differentiation of wine, brewing, bakery and sake strains of Saccharomyces species Genetic analysis of S. cerevisiae wine strains Identification of pathogenic Candida species, subspecies of C. albicans and C. dublinensis Intraspecific variability among A. pullulans

Azumi and Goto-Yamamoto (2000)

Differentiation of S. cerevisiae, S. pastorianus, S. bayanus, S. willianus Differentiation of C. neoformans, C. albidus, C. laurentii and R. rubra species, and strains of C. neoformans Differentiating S. cerevisiae and Z. bailii Characterisation of wine yeasts; R. mucilaginosa, S. cerevisiae, S. exiguus, P. membranifaciens, P. anomala, T. delbrueckii, C. vini strains Distinguish species within genus Saccharomyces

Lieckfeld et al. (1993)

Gallego et al. (2005) Borst et al. (2003), Ball et al. (2004) De Curtis et al. (2004)

RAPD and micro/minisatellites (GTG)5 (CAG)5 and M13 (GTG)5, (GACA)4 and phage M13 core sequence Primers 15, 18, 20, 21 Preliminary screening of primers

Decamer 1; ACG GTG TTG G Decamer 2; TGC CGA GCT G Decamer 3; GGG TAA CGC C M13

Baleiras Couto et al. (1994) Quesada and Cenis (1995)

Molnár et al. (1995) Lopandic et al. (1996) Herzberg et al. (2002) Prillinger et al. (1999)

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Distinguish species within genus Metschnikowia Yeast species isolated from floral nectaries Identify yeast species from cheese; D. hansenii, S. cerevisiae, I. orientalis, K. marxianus, K. lactis, Y. lipolytica, C. catenulata, G. candidum

Meyer et al. (1993)

Evaluation of Molecular Methods for the Analysis of Yeasts in Foods

Table 4. Application of PCR-based methods for strain and species differentiation of yeasts associated with foods and beverages Primers Application References

Continued

82 Ai Lin Beh et al.

Table 4. Application of PCR-based methods for strain and species differentiation of yeasts associated with foods and beverages—cont’d Primers Application References Decamer 1; ACG GTG TTG G Decamer 2; TGC CGA GCT G Decamer 3; TGC AGC GTG G Decamer 4; GGG TAA CGC C M13 Yeast species from dairy products; S. cerevisiae, Andrighetto et al. (2000) K. marxianus, K. lactis, D. hansenii, Y. lipolytica, T. delbrueckii RF2 Yeast species from sourdough products Forshino et al. (2004) M13V universal Yeast species from Greek sourdough; Paramithiotis et al. (2000) P. membranifaciens, S. cerevisiae, Y. lipolytica M13 Yeast species from artisanal Fiore Sado cheese; Fadda et al. (2004) C. zeylanoides, D. hansenii, K. lactis, C. lambica, G. candidum Vasdinyei and Deak (2003) Strain typing of D. hansenii and G. candidum Primers 24, 28, OPA11 Baleiras Couto et al. (1996a) Discrimination of S. cerevisiae strains (GAC)5, (GTG)5 Baleiras Couto et al. (1996b) (GTG)5 and (CAG)5 Differentiation of strains of Z. bailii and Z. bisporus Caruso et al. (2002) Separation of S. cerevisiae from K. apiculata Capece et al. (2003) Separation of C. stellata, M. pulcherrima, K. apiculata and Schiz. pombe (GTG)5, (CAG)5 and M13 Genotyping the R. glutinis complex Gadanho and Sampaio (2002) Primer P24, (GTG)5 and (GAC)5 Pina et al. (2005) Typing of P. galeiformis strains in orange juice production (GTG)5 (ATG)5 and M13 Cadez et al. (2002) Differentiation of Hanseniaspora species OPA03, OPA 18

REPIR1, REP2I Intron splice site primer EI1 Intron splice primers

C. boidinii, C. mesenterica, C. sake, C. stellata, D. anomala, D. bruxellensis, H. uvarum, I. terricola, S. ludwigii, Schiz. pombe, T. debrueckii, Z. bailii, S. bayanus, S. cerevisiae from wine

Hierro et al. (2004)

Differentiation of commercial wine S. cerevisiae strains. Differentiation of non-Saccharomyces wine species and strains; S. cerevisiae, S. bayanus, T. delbrueckeii, I. orientalis, H. uvarum, H. guillermondii, M. pulcherrima, P. fermentans, P. membranaefaciens

de Barros Lopes et al. (1996, 1998)

EI1, EI2, LA1, LA2

S. cerevisiae and sensu stricto strains Primers δ1 and δ2

Primers δ12 and δ2

Introns in COX1 mitochondrial gene

Differentiation of S. cerevisiae strains, S. douglassi, S. chevalierii, S. bayanus

Ness et al. (1993) Lavallée et al. (1994), Fernández-Espinar et al. (2001), Schuller et al. (2004)

Identification/authentication of commercial wine S. cerevisiae strains Characterisation of wild S. cerevisiae strains from grapes and wine fermentations Differentiating S. cerevisiae strains from sourdough Genetic relatedness between clinical and food S. cerevisiae strains Monitor wine starter S. cerevisiae strains during fermentation

Versavaud et al. (1995), Cappello et al. (2004), Demuyter et al. (2004) Pulvirenti et al. (2001) de Llanos et al. (2004) López et al. (2003)

Evaluation of Molecular Methods for the Analysis of Yeasts in Foods

ERIC1R, ERIC2

Continued

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Table 4. Application of PCR-based methods for strain and species differentiation of yeasts associated with foods and beverages—cont’d Primers Application References

Microsatellite markers/Sequence-Tagged Site markers (ScTAT1) chromosome XIII Gallego et al. (1998) Differentiation of S. cerevisiae strains Locus SCYOR267, Locus SC8132X, Differentiation of industrial S. cerevisiae wine strains González Techera et al. (2001) Locus SCPTSY7 Locus SC8132X Monitoring the populations of S. cerevisiae Howell et al. (2004) strains during grape juice fermentation Multiplex 1; Loci ScAAT2, ScAAT3, ScAAT5 Pérez et al. (2001), Gallego et al. (2005) Differentiation of S. cerevisiae isolates from spontaneous wine fermentations Multiplex 2; Loci ScAAT1, ScAAT4, ScAAT6 Differentiation of commercial S. cerevisiae Schuller et al. (2004) wine strains Minisatellite core sequence (wild type phage) GAG GGT GGC GGT TCT; M13V universal; GTT TCC CCA GTC ACG AC; Phage M13 core sequence; GAG GGT GGX GGX TCT

Ai Lin Beh et al.

Evaluation of Molecular Methods for the Analysis of Yeasts in Foods

2.5.

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PCR-based Fingerprinting

PCR technology has provided new opportunities for developing faster, more convenient methods for typing yeasts and fungi. Two of the first methods developed were Random Amplified Polymorphic DNA (RAPD) analysis and Amplified Fragment Length Polymorphism (AFLP) analysis (Baleiras Couto et al., 1994, 1995, 1996a; Vos et al., 1995; van der Vossen et al., 2003). These methods have the capability of analyzing an extensive portion of the genome, and reveal polymorphisms that differentiate at both the species and strain levels (Table 4). In the case of RAPD, DNA template is subject to PCR amplification with single, short primers (10-15bp of random/arbitary sequences) that hybridize to a set of arbitary loci in the genome. Some primers used for this purpose are listed in van der Vossen et al. (2003). PCR cycles are performed under conditions of low stringency. The amplicons produced are separated on gel electrophoresis and give profiles that fingerprint the strain or species. AFLP is a variation of RAPD. The approach firstly involves digestion of genomic DNA with two restriction nucleases (usually EcoRI and MseI). The fragments are then ligated with end-specific adapters, followed by two successive rounds of PCR. Pre-selective PCR amplifies fragments using primers complimentary to the adapter sequences. A second, selective PCR is performed with primers containing additional nucleotide bases at the 3′ end (selective bases are user defined). The resulting products are resolved by gel electrophoresis or by capillary electrophoresis. There are extra steps involved in AFLP, but this method can potentially generate more extensive information from a single restriction/ligation reaction than other PCR strategies (de Barros Lopes et al., 1999; Lopandic et al., 2005). Both RAPD and AFLP analyses gave excellent strain and species differentiation. However, the main hurdles to routine application are the need for stringent standardisation of conditions in order to obtain reproducible data, and the workload involved. Micro- and minisatellites are short repeat motifs of about 15-30 and 2-10bp, respectively. Primers targeting these sequences are used in PCR assays to generate an array of amplicons, the profile or “fingerprint” of which reflects the polymorphism of these regions and the distance between them. Some commonly used primers for food yeasts are (GTG)5, (GAC)5 and M13 phage core sequences. This method has had good success in differentiating strains of several food spoilage and wine yeasts (Baleiras Couto, et al., 1996b), and for differentiating species of yeasts from dairy sources (Prillinger et al., 1999) (Table 4).

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Intron splice sequences are also known for their polymorphism and have been examined as sites that could give strain differentiation. De Barros Lopes et al. (1996, 1998) reported a PCR method for the analysis of these sites. The method was simple, quick (several hours), robust, reproducible and gave profiles enabling the differentiation of winemaking strains of Saccharomyces. Building on this concept, López et al. (2002) developed a multiplex PCR assay based on introns of the COX1 mitochondrial gene. The assay, which was simple and fast (8 hours), gave good differentiation of wine strains of Saccharomyces. Other repetitive elements that are targeted in PCRmediated fingerprinting techniques include the delta repeat, the repetitive extragenic palindromic (REP) and enterobacterial repetitive intergenic consensus (ERIC) sequences (Hierro et al., 2004). The application of these PCR methods to food and beverage yeasts (Table 4) generally gives good species and strain characterisation. However, the level of resolution achieved is greatly influenced by the choice of primers and the taxa under study.

3.

MOLECULAR STRATEGIES FOR MONITORING YEAST COMMUNITIES IN FOODS AND BEVERAGES

The diversity of yeast species associated with foods and beverages is usually determined by culturing homogenates of the product on plates of agar media (Fleet, 1992; Deak, 2003). Yeast colonies are then isolated and identified. As mentioned in previous sections, molecular methods have now found widespread application in identifying these isolates. New, culture-independent methods based on PCR-denaturing gel gradient electrophoresis (DGGE) and PCR-temperature gradient gel electrophoresis (TGGE) are now being used to determine the ecological profile of yeasts in foods and beverages (Muyzer and Smalla, 1998). The basic strategy is outlined in Figure 2. Total DNA is extracted from samples of the product. Using universal fungal primers (or genus-specific primers), yeast ribosomal DNA within the extract is specifically amplified by PCR. Generally, the D1/D2 domain of the 26S subunit is targeted, but other regions such as the 18S subunit may be used. The amplicons produced by PCR are next separated by either DGGE or TGGE, which resolve the different DNA amplicons on the basis of their sequence/melting domains. DGGE uses a polyacrylamide gel containing a linear gradient of denaturant (mix-

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Food / Beverage Extract and purify DNA/RNA PCR-amplification of yeast rDNA DGGE/TGGE separation of DNA amplicons DNA bands correspond to different species

Isolate and sequence bands to give species identity

Figure 2. A culture-independent approach for determining the yeast ecology of foods and beverages using PCR-DGGE/TGGE analyses

ture of urea and formamide), while TGGE uses a gradient of temperature to denature the strands of DNA amplicons. Usually, each DNA band found in the gel corresponds to a yeast species. The band is excised from the gel and sequenced to give the species identity. Thus, a profile of the species associated with the ecosystem is obtained, without the need for agar culture. It is believed that this molecular approach overcomes the bias of culture methods, and reveals species that might fail to produce colonies on agar media. Consequently, it is considered that a more accurate representation of the diversity of yeast species in the food product is obtained (Giraffa, 2004). Table 5 lists a range of studies where PCR-DGGE/TGGE have been applied to the analysis of food and beverage yeasts. Generally, there has been good agreement between yeast species detected in foods and beverages by PCR-DGGE/TGGE and culture on agar media, but some discrepancies have been noted. In some cases, yeasts were recovered by DGGE/TGGE analyses but not by culture. These observations have led to suggestions that viable but nonculturable yeasts may be present (Mills et al., 2002; Meroth et al., 2003; Masoud et al., 2004; Prakitchaiwattana et al., 2004; Nielsen et al., 2005). However, DNA from non-viable yeast cells or DNA released from autolysed yeast cells could account for these findings, and caution is needed when interpreting the DGGE/TGGE data. To address this limitation, Mills et al. (2002) and Cocolin and Mills

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Table 5. Application of PCR-denaturation gradient gel electrophoresis and PCR-temperature gradient gel electrophoresis to ecological studies of yeasts in foods and beverages. Method: region/primers Application References PCR-DGGE Succession of yeast species in wine fermentations; Nested PCR; NL1-NL4, NL1gc-LS2; 26S rDNA model wine fermentation; Cocolin et al. (2000) commercial Dolce wine fermentations; Cocolin et al. (2001) continuous wine fermentation; Cocolin et al. (2002a) PCR-DGGE and RT PCR-DGGE Yeast species in Botrytis-affected wine fermentations Mills et al. (2002) 26S rDNA/rRNA; nested PCR; Inhibition of wine yeast species by sulphur dioxide Cocolin and Mills (2003) NL1-NL4, NL1gc-LS2 PCR-TGGE Discrimination of wine yeast species Hernán-Gómez et al. (2000) 18S rDNA; YUNIV1gc-YUNIV3 Diversity of yeast species in wine fermentations Fernández-Gonzáles et al. (2001) PCR-DGGE Yeast species in sourdough starters mixtures, rye Meroth et al. (2003) flour and sourdough samples 26S rDNA; U1gc-U2 PCR-DGGE Yeast species on wine grapes Prakitchaiwattana et al. (2004) 26S rDNA; nested PCR; NL1-NL4, NL1gc-LS2

18S rDNA; nested PCR; NS1-NS8, NS1gc-NS2+10 PCR-DGGE 26S rDNA; NL1gc-LS2 PCR-DGGE 26S rDNA; nested PCR; NL1-NL4, NL1gc-LS2; or NL1-LS2, NL1gc-LS2 PCR-DGGE 26S rDNA; NL1gc-LS2 PCR-DGGE ITS2; Schafgc-Schar (Saccharomyces sensustricto specific) PCR-TGGE ITS2; Schafgc-Schar

Phyllospheric yeast species and fungicide treatments on russetting of Elstar apples

Gildemacher et al. (2004)

Yeast species in raw milk

Cocolin et al. (2002b)

Yeast species involved in coffee fermentation

Masoud et al. (2004)

Yeast populations associated with Ghanaian cocoa fermentations Differentiation of Saccharomyces sensu strictu strains; S. cerevisiae, S. paradoxus and S. bayanus/S. pastorianus

Nielsen et al. (2005)

S. cerevisiae and S. paradoxus in wine fermentation

Manzano et al. (2004)

Manzano et al. (2005)

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(2003) have proposed the use of RT-PCR protocols that target RNA rather than DNA templates from the food or beverages. In several studies, culture methods have revealed the presence of yeast species not found by PCR-DGGE (Mills et al., 2002; Masoud et al., 2004; Prakitchaiwattana et al., 2004). Such yeasts were present at populations at less than 102-103 CFU/ml or g of product and this is considered to be the lower limit of detection by PCR-DGGE. Nevertheless, there are reports where yeast species present at 104-105 CFU/ml or g product have not been detected by PCR-DGGE. This can occur when there is a mixture of different yeast species in the sample, with one species being numerically present at populations 1001000 times more than other species. The DNA of the dominant species may bind primers more favorably, resulting in weaker amplification of the minority species. Also, when universal primers are used, there may be stronger hybridization of the primers with the DNA of some species over others (Mills et al., 2002; Prakitchaiwattana et al., 2004; Nielsen et al., 2005). While the relative mobility of DNA bands on DGGE/TGGE gels can discriminate between closely related species (e.g. those in Saccharomyces sensu stricto, Manzano et al., 2004), exceptions can occur. In some cases, different species have produced bands with similar mobilities (Hernán-Gómez et al., 2000; Gildemacher et al., 2004). There are several reports where multiple bands have been found for the one species (eg. two bands for Candida sp. (Mills et al., 2002), three bands for Pichia kluyveri (Masoud et al., 2004), and several bands for Metschnikowia pulcherrima (Prakitichaiwattana et al., 2004). The reasons for multiple banding within the one species are not well understood, but may reflect artefacts of PCR using primers with GC clamps and DNA denaturation kinetics during electrophoresis. Multiple bands could arise from nucleotide variations among multiple rDNA copies within a single strain, or could also indicate the presence of different strains within a species. To address these anomalies, it is good practice therefore, to isolate individual bands from DGGE/TGGE gels and confirm their identity by sequencing. Finally, food samples often contain large amounts of DNA from plants, animals and other microbial groups (eg. bacteria and filamentous fungi) that have the potential to interfere with the specific PCRamplification of yeast DNA and compromise the reliability and quality of the data obtained by DGGE/TGGE. For example, we have experienced particular difficulty in detecting yeasts in mould ripened cheeses with PCR-DGGE and PCR-TGGE because of the large

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amounts of fungal (Penicillium) DNA that occur in the cheese extracts (Nurlinawati, Cox and Fleet, unpublished data).

4.

FACTORS AFFECTING PERFORMANCE OF MOLECULAR METHODS FOR THE ANALYSIS OF YEASTS

As mentioned previously, molecular methods need to meet certain performance and practical criteria before they will gain acceptance for routine use in the quality assurance laboratories of the food and beverage industries. First, they will need to give accurate, reliable and reproducible data at the appropriate levels of sensitivity and selectivity, and meet the appropriate tolerances for false positive and false negative results. Second, they will need to be simple, convenient and inexpensive to use, and give results relatively quickly (Cox and Fleet, 2003). PCR assays form the basis of most molecular methods used to analyse yeasts. Consequently, it is important to identify and understand the various factors which affect the performance of this technology. The principles of PCR can be found in many text books (eg. McPherson and Møller, 2000). More specific discussions of its application in microbiological analysis are given by Bridge et al. (1998), Hill and Jinneman (2000), Sachse and Frey (2003) and Cox and Fleet (2003). The following sections highlight some of the conceptual and practical variables that affect its performance as applied to the analysis of food and beverage yeasts. More general discussions of these factors are given by Edel (1998), Hill and Jinneman (2000), Sachse (2003), Radström et al. (2003), Bretagne (2003) and Lübeck and Hoofar (2003). PCR assay involves the following operations; (i) sample preparation (ii) extraction and preparation of DNA (iii) amplification of DNA by PCR (iv) detection of PCR products and (v) processing and interpretation of the data.

4.1.

Sample Preparation and DNA Extraction

For many applications, the sample is a pure culture of a yeast isolate. Consequently, sampling is not an issue provided that the culture has proven purity. Nevertheless, the culture needs to be grown to provide biomass for DNA extraction. Variables here include the

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culture medium and time of incubation, and these have not been given proper consideration in previous literature. Carry over of media ingredients could inhibit the PCR assay (Rossen et al., 1992). The physiological age of the yeast cells (e.g. exponential, stationary phase, autolysing, dead) at the time of assay can affect the efficiency of DNA extraction and the quality of DNA template for PCR. Depending on culture age, various cell proteins, for example, may interact with the genomic DNA, thereby affecting primer annealing to the template, or they can affect the activity of the DNA polymerase (de Barros Lopes et al., 1996). Some basidiomycetous yeasts may have tougher cell walls than ascomycetous yeasts and require more vigorous procedures for equivalent DNA extraction (Prakitchaiwattana et al., 2004). The purity and concentration of template DNA become more critical when analysing yeast cells associated with food or beverage matrices, as for example, in studies using DGGE or TGGE (Table 5). It is well known that plant polysaccharides, humic components and other polyphenolic materials can co-purify with DNA and inhibit the PCR reaction (Wilson, 1997; Marshall et al., 2003). The relative ratio of yeast DNA to other DNA species (e.g. that from filamentous fungi, bacteria, plants) is another factor that is not properly understood in the performance of PCR assays. The design of primers would be very important here to minimise or prevent their binding to non-target DNA. Many “in-house” methods have been described to extract and purify DNA from yeast cells, whether they be biomass originating from a pure culture or biomass extracted directly from the food matrix. The methods include freezing and boiling of cells, mechanical disruption by shaking with glass beads or zirconium, digestion with lytic enzymes and extraction with chemical solvents (Hill and Jinneman, 2000; Haugland et al., 2002). Ideally, this “front-end” part of the analytical process needs to be simple and convenient, but yield template DNA that will perform satisfactorily in PCR assays and give the required detection sensitivity. Critical evaluation and some degree of standardization of these methods are needed.

4.2.

PCR Amplification of Template DNA

PCR assays are enzymatic reactions. Accordingly, their performance and progress follow the basic principles of enzymology. The substrates are the template DNA, oligonucleotide primers and equimolar amounts of each nucleotide base; ATP, GTP, TTP and CTP, the enzyme to catalyze the reaction is DNA polymerase, and the product

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is newly synthesized DNA. Like all enzymatic processes, the reaction is highly specific, and its kinetics are determined by factors such as pH, temperature, concentration of reactants, requirements for cofactors and the presence of any inhibitors. Magnesium ions are critical co-factors, the concentration of which determines the specificity, efficiency and fidelity of the reaction. Two factors make PCR more complex than other enzyme reactions. First, temperature control requires cyclic variation according to the following program; starting at 91-97’C for the denaturation of double stranded DNA; decreasing to 40-65’C for primer annealing to single strands of template DNA; and increasing to 68-74’C for DNA strand extension by DNA polymerase. About 30-40 cycles are conducted. Second, the DNA product also becomes an enzyme substrate as the reaction progresses. The great diversity of PCR applications also introduces specific variables that require understanding and optimization. In particular, these are the length and sequence of primers, the amount of template DNA, the amount of non-target/background DNA, and carry-over material from the sample matrix that could inhibit primer annealing and DNA polymerase activity. Table 6 summarizes some of the key variables that affect the performance of PCR assays. Because of the broad range of PCR applications, it is difficult to prescribe one set of optimum conditions. Consequently, optimisation must be done on the basis of each application. The key aims of optimisation are to increase diagnostic specificity and diagnostic sensitivity (detection limit). Some good discussions of these variables can be found in Edel (1998), McPherson and Møller (2000), Sachse (2003), Radström et al. (2003), Bretagne (2003) and Lübeck and Hoofar (2003). Wilson (1997) has reviewed various factors that inhibit and facilitate PCR. The conditions of electrophoresis used to separate and detect the DNA amplicons represent another suite of variables that need

Table 6. Factors affecting DNA amplification by PCR 1. Primer to template ratio 2. Efficiency of primer annealing 3. Enzyme to template ratio 4. Length and sequence of primers 5. Concentration of non-target DNA 6. Inhibitors from sample matrix 7. Temperature cycling protocol 8. Source of DNA polymerase 9. Reaction facilitators

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to be optimised and managed. Variables, here include the gelling agent (agarose or polyacrylamide) and its concentration, running time, temperature and voltage, and composition of buffer (Andrews, 1986; Hames, 1998). The extent of cross-linking and pore size in polyacrylamide gels affected the resolution of DNA bands and detection of yeast species on grapes by PCR-DGGE (Prakitchaiwattana et al., 2004).

5.

STANDARDIZATION OF MOLECULAR METHODS FOR ANALYSIS FOR YEASTS IN FOODS AND BEVERAGES

The commercial significance of yeast in foods and beverages has major implications in national and international trade (Fleet, 2001). Companies trading in foods and beverages will usually have contractual arrangements that specify criteria for the presence of yeasts. In this context, molecular methods used for yeast analyses will need to meet the rigours of legal or forensic scrutiny. They will need some form of standardization and international acceptance. While there have been major advances in the harmonization and standardization of cultural methods for the analysis of microorganisms in food and beverages (Food Control, 1996; Scotter et al., 2001; Langton et al., 2002), this need remains a challenge for molecular methods. The lack of standardization and validation for molecular analyses of microorganisms in foods and beverages is well recognized and strategies are being developed to address this need, especially for bacteria of public health significance (Schafer et al., 2001; Lübeck and Hoofar, 2003; Lübeck et al., 2003). Hoofar and Cook (2003) and Malorny et al. (2003) have outlined the principles and protocols for achieving this goal, based upon the FOOD-PCR project of the European Commission (http://www.PCR.dk). These initiatives equally apply to the molecular analyses of yeasts in foods and provide a framework upon which to develop similar projects for yeasts and other fungi. The first stage is to define the specific application to be evaluated. This is followed by evaluating and defining the conditions of the assay (e.g. sample treatment and DNA extraction, primer selection, PCR conditions, detection limit), developing positive and negative control assays, selecting procedures for data analysis, and, finally, developing protocols for “in-house” and inter-laboratory validation trials. From these evaluations, consensus and standardization should

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emerge. Leuschner et al. (2004) reported the results of an interlaboratory evaluation of a PCR method for the detection and identification of probiotic strains of S. cerevisiae in animal feed. Good agreement (but not 100%) was obtained between laboratories and an “official” method for analysing animal feed for these yeasts was proposed.

6.

REFERENCES

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scriptase sequencing of plasma samples from heavily treated patients. J. Clin. Microbiol. 39:1522-1529. Schuller, D., Valero, E., Dequin, S., and Caral, M., 2004, Survey of molecular methods for the typing of wine yeast strains, FEMS Microbiol. Lett. 231:19-26. Scorzetti, G., Fell, J. W., Fonseca, A., and Statzell-Tallman, A., 2002, Systematics of basidiomycetous yeasts: a comparison of large subunit D1/D2 and internal transcribed spacer rDNA regions, FEMS Yeast Res. 2:495-517. Scotter, S. L., Langton, S., Lombard, B., Lahellec, C., Schultern, S., Nagelkerke, N., in’t Veld, P. H., and Rollier, P., 2001, Validation of ISO method 11290 Part 2. Enumeration of Listeria monocytogenes in foods, Int. J. Food Microbiol. 70: 121-129. Sipiczki, M., 2002, Taxonomic and physiological diversity of Saccharomyces bayanus in: Biodiversity and Biotechnology of Wine Yeasts. M. Ciani, ed., Research Signpost, Kerala, India, pp. 53-69. Smole Mozina, S., Dlauchy, D., Deak, T., and Raspor, P., 1997, Identification of Saccharomyces sensu stricto and Torulaspora yeasts by PCR ribotyping, Lett. Appl. Microbiol. 24:311-315. Spear, R. N., Li, S., Nordheim, E. V., and Andrews, J. H., 1999, Quantitative imaging and statistical analysis of fluorescence in situ hybridization (FISH) of Aureobasidium pullulans, J. Microbiol. Methods 35:101-110. Stender, H., HHKurtzman, C., Hyldig-Nielsen, J. J., Sorensen, D., Broomer, A., Oliveira, K., Perry-O’Keefe, H., Sage, A., Young, B., and Coull, J., 2001, Identification of Dekkera bruxellensis (Brettanomyces) from wine by fluorescence in situ hybridization using peptide nucleic acid probes, Appl. Environ. Microbiol. 67:938–941. Stratford, M., Bond, C. J., James, S. A., Roberts, I. N., and Steels, H., 2002, Candida davenportii sp. nov., a potential soft-drinks spoilage yeast isolated from a wasp, Int. J. Syst. Evol. Microbiol. 52:1369-1375. Suzuki, M., and Nakase, T., 1999, A phylogenetic study of ubiquinone Q-8 species of the genera Candida, Pichia, and Citeromyces based on 18S ribosomal DNA sequence divergence, J. Gen. Appl. Microbiol. 45:239-246. Torriani S., Zapparoli G., Malacrino P., Suzzi G., and Dellaglio, F., 2004, Rapid identification and differentiation of Saccharomyces cerevisiae, Saccharomyces bayanus and their hybrids by multiplex PCR, Lett. Appl. Microbiol. 38:239-244. Valente, P., Ramos, J. P., and Leoncini, O., 1999, Sequencing as a tool in yeast molecular taxonomy, Can. J. Microbiol. 45:949-958. van Keulen, H., Lindmark, D. G., Zeman, K. E., and Gerlosky, W., 2003, Yeasts present during spontaneous fermentation of Lake Erie Chardonnay, Pinot Gris and Riesling, Antonie van Leeuwenhoek 83:149-154. van der Aa Kühle, A., Jespersen, L., Glover, R. L. F., Diawara, B., and Jakobsen, M., 2001, Identification and characterization of Saccharomyces strains from West African sorghum beer, Yeast 18:1069-1079. van der Aa Kühle, A., and Jespersen, L., 2003, The taxonomic position of Saccharomyces boulardii as evaluated by sequence analysis of the D1/D2 domain of 26S rDNA, the ITS1-5.8S rDNA-ITS2 region and the mitochondrial cytochrome-c oxidase II gene, System. Appl. Micobiol. 26:564-571. van der Vossen, J. M. B. M., Rahaoui, H., de Nijs, M. W. C., and Hartog, B. J., 2003, PCR methods for tracing and detection of yeasts in the food chain, in: Yeasts in Food, Beneficial and Detrimental Aspects, T. Boekhout and V. Robert, eds, Behr’sVerlag, Hamburg, pp. 123-138.

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Vasdinyei R., and Deák, T., 2003, Characterization of yeast isolates originating from Hungarian dairy products using traditional and molecular identification techniques, Int. J. Food Microbiol. 86:123-130. Versavaud A., Courcoux P., Roulland C., Dulau L., and Hallet J. N., 1995, Genetic diversity and geographical distribution of wild Saccharomyces cerevisiae strains from the wine-producing area of Charentes, France. Appl. Environ. Microbiol. 61:3521-3529. Villa-Carvajal, M., Coque, J. J. R., Álvarez-Rodríguez, M. L., Uruburu, F., and Belloch, C., 2004, Polyphasic identification of yeasts isolated from bark of cork oak during the manufacturing process of cork stoppers, FEMS Yeast Res. 4:745-750. Vos, P., Hogers, R., Bleeker, M., Reijans, M., van de Lee, T., Hornes, M., Frijters, A., Pot, J., Peleman, J., Kuiper, M., and Zabeau, M., 1995, AFLP: a new technique for DNA fingerprinting, Nucleic Acids Res. 23:4407-4414. White, T. J., Bruns, T., Lee, S., and Taylor, J., 1990, Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenentics, in: PCR Protocols: A Guide for Methods and Applications. M. A. Innis, D. H. Gelfand, J. J. Sninsky and T. J. White, eds, Academic Press, San Diego, pp. 315-352. Wilson, I. G., 1997, Inhibition and facilitation of nucleic acid amplification. Appl. Environ. Microbiol. 63:3741-3751.

STANDARDIZATION OF METHODS FOR DETECTING HEAT RESISTANT FUNGI Jos Houbraken and Robert A. Samson*

1.

INTRODUCTION

Heat resistant fungi can be defined as those capable of surviving temperatures at or above 75°C for 30 or more minutes. The fungal structures which can survive these temperatures are ascospores, and sometimes chlamydospores, thick walled hyphae or sclerotia (Scholte et al., 2000). During the last three years, spoilage incidents involving heat resistant fungi occurred increasingly in various products examined in our laboratory. Paecilomyces variotii, Fusarium oxysporum, Byssochlamys fulva, B. nivea, Talaromyces trachyspermus and Neosartorya species were often encountered in pasteurized fruit, dairy products and soft drinks. A questionnaire sent to many laboratories showed that inappropriate methods were used for the detection of heat resistant fungi, or that sometimes there was no special protocol at all. The use of inappropriate media, such as Sabouraud agar, wrong incubation conditions and the analysis of inadequately sized samples were often encountered. In addition, accurate identification of the isolated colonies to species level often was not performed. In the literature many methods are described for the detection of heat resistant fungi (Murdock and Hatcher, 1978; Beuchat and Rice, 1979; Beuchat and Pitt, 1992). Beuchat and Pitt (1992) described two methods: the Petri dish method or plating method and the direct incubation method. In the first method, test tubes are used for the heating of the sample. Subsequently, the sample is poured into large Petri * Centraalbureau voor Schimmelcultures, PO Box 85167, 3508 AD, Utrecht, The Netherlands. Correspondence to: [email protected]

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dishes (diameter 14 cm) and agar media is added. In the second method, flat-sided bottles are used for heating and these bottles are incubated directly at 30°C. The direct incubation method has the advantage that contamination from aerial spores is minimized; the disadvantage of this method is that the colonies growing in the bottle have to be transferred onto agar media for identification. During recent years a great variety of products have been investigated in our laboratory for the presence of heat resistant fungi. Some products were liquids (juices, colourants), some with a high viscosity (fruit concentrates), or solid products such as soil, pectin, liquorice, strawberries and cardboard. For several of these products the recommended methods are not suitable. A modified detection method using Stomacher bags is therefore proposed here.

2.

METHOD FOR EXAMINATION FOR HEAT RESISTANT MOULDS

2.1.

Main Modifications

The modified method proposed here is based onto the protocol of Pitt and Hocking (1997). The major modification is the use of Stomacher bags for the heating step which is the important step in the isolation of heat resistant fungi. The product can be easily homogenized in Stomacher bags, with a low risk of aerial contamination. The ascospores of many species of heat resistant fungi require heat activation before they will germinate (Katan, 1985; Lingappa and Sussman, 1959). The heat treatment also inactivates vegetative cells of fungi and bacteria, as well as less heat resistant spores (Beuchat and Pitt, 1992). Some protocols require that bottles be used for the heating of the sample. If bottles that are circular in cross section rather than flat-sided are used, the heat penetration into the sample is particularly slow. When a Stomacher bag containing 250 ml of sample is sealed half way along its length, the thickness of the bag with the sample will be little more than 1 cm, providing much faster heat penetration than in a 250 ml bottle.

2.2.

Description of the Modified Method

Because the concentration of the heat resistant fungi in a sample is normally very low, the analysis of a large amount of sample is recom-

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mended. At least 100 g of sample should be examined (Samson et al., 2000). Homogenize the sample before beginning the analysis. Transfer 100 g of sample into a sterile Stomacher bag. Add sterile water (150 g) to the sample and homogenize using a Stomacher for 2 to 4 min. If the sample is likely to contain a higher concentration of heat resistant fungi (e.g. a soil sample), or the sample cannot be homogenized in 150 grams of water (e.g. solid ingredients such as pectin), then a smaller amount of sample can be used. After the homogenization step, the Stomacher bag should be sealed about half way along its length using a heat-sealer, ensuring that no air bubbles are present. After checking the bag for leakage, heat treat the Stomacher bag for 30 min at 75°C in a water bath (preferably one with the capability of shaking or stirring). The water-bath should be at 75°C before the sample is introduced. The sample should be placed in a horizontal position, totally submerged in the water. After the heat treatment, cool the samples to approximately 55°C. Aseptically transfer the contents of the Stomacher bag to a Schott bottle, or similar, (500 ml) with 250 ml melted double strength MEA containing chloramphenicol (200 mg/l, Oxoid) tempered to 55°C. Mix thoroughly and distribute the agar and sample mixture into seven large plastic Petri dishes (diameter 14.5 cm). Place the Petri dishes into a polyethylene bag to prevent drying and incubate in an upright position at 30°C in darkness. The general procedure is illustrated in Figure 1.

2.3.

Incubation

Many protocols require plates to be incubated for at least 30 days. This period is too long for quality control in the food and beverage industry. In our experience, heat-resistant fungi will usually form colonies after 5 days and mature within 14 days incubation at 30°C, so check the Petri dishes for the presence of colonies after 7 and 14 days. Subculture if necessary, and identify all colonies using standard methods (Pitt and Hocking, 1997; Samson et al., 2000). In some cases a prolonged incubation period is necessary for the identification of the fungus. An overview of the incubation time needed for some particular species is given in Figure 2.

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Homogenize Stomacher bag

H2O

and seal

Sample 1. Liquid samples: 100 ml + 150 ml water 2. Pectin: 12.5 g + 230 ml water 3. Fruit (concentrate): 100 g + 150 ml water 4. Solid samples (eg cardboard, powdery ingredients, soil): 25 g + 225 ml water

1. Incubate for 14 d. at 30°C 2. Check every 7 days

Heat treatment, 30 min 75°C

Mix throughly and disperse agar/product mass into approx. 7-8 petri-dishes (diam. 14.5 cm)

1. Cool the sample rapidly 2. Mix the sample with 250 ml handwarm double strength MEA agar + chloramphenicol

Figure 1. Modified method for the detection of heat resistant fungi Talaromyces trachyspermus Eupenicillium species, Monascus ruber T. macrosporus, Hamigera sp. Byssochlamys sp., P. variotii s.l., Neosartorya sp. Air contamination or under pasteurization (Rhizopus, Humicola, Penicillia, A. niger / flavus (also sclerotial isolates) 0

2

4

6

8

10

12

14 incubation time (days)

Figure 2. Overview of the incubation time needed to allow growth and development for particular species of heat resistant fungi

3.

REFERENCES

Beuchat, L. R., and Rice, S. L., 1979, Byssochlamys spp. and their importance in processed fruit syrups, Trans. Br. Mycol. Soc. 68:65-71. Beuchat, L. R., and Pitt, J. I., 1992, Detection and enumeration of heat-resistant molds, in: Compendium for the Microbiological Examination of Foods, C. Vanderzant and D. F. Splittstoesser, eds, American Public Health Association, Washington D. C., pp. 251-263.

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Katan, T., 1985, Heat activation of dormant ascospores of Talaromyces flavus, Trans. Br. Mycol. Soc. 84:748-750. Lingappa, Y., and Sussman, A. S., 1959, Changes in the heat resistance of ascospores of Neurospora upon germination, Am. J. Botany 49:671-678. Murdock, D. I., and Hatcher, W. S., 1978, A simple method to screen fruit juices and concentrates for heat-resistant mold, J. Food. Prot. 41:254-256. Pitt, J. I., and Hocking, A. D., 1997, Fungi and Food Spoilage, 2nd edition, Blackie Academic and Professional, London. Samson, R. A., Hoekstra, E. S., Lund, F., Filtenborg, O., and Frisvad, J. C., 2000, Methods for the detection, isolation and characterization of food-borne fungi, in: Introduction to Food- and Airborne Fungi, 6th edition, R. A. Samson, E. S. Hoekstra, J. C. Frisvad and O. Filtenborg, eds, Centraalbureau voor Schimmelcultures, Utrecht, pp. 283-297. Scholte, R. P. M., Samson, R. A. and Dijksterhuis, J., 2000, Spoilage fungi in the industrial processing of food, in: Introduction to Food- and Airborne Fungi, 6th edition, R. A. Samson, E. S. Hoekstra, J. C. Frisvad, and O. Filtenborg, eds, Centraalbureau voor Schimmelcultures, Utrecht, pp. 339-356.

Section 3. Physiology and ecology of mycotoxigenic fungi Ecophysiology of fumonisin producers in Fusarium section Liseola Vicente Sanchis, Sonia Marín, Naresh Magan, and Antonio J. Ramos Ecophysiology of Fusarium culmorum and mycotoxin production Naresh Magan, Russell Hope and David Aldred Food-borne fungi in fruit and cereals and their production of mycotoxins Birgitte Andersen and Ulf Thrane Black Aspergillus species in Australian vineyards: from soil to ochratoxin A in wine Su-lin L. Leong, Ailsa D. Hocking, John I. Pitt, Benozir A. Kazi, Robert W. Emmett and Eileen S. Scott Ochratoxin A producing fungi from Spanish vineyards Marta Bau, M. Rosa Bragulat, M. Lourdes Abarca, Santiago Minguez, and F. Javier Cabañes Fungi producing ochratoxin in dried fruits Beatriz T. Iamanaka, Marta H. Taniwaki, E. Vicente and Hilary C. Menezes An update on ochratoxigenic fungi and ochratoxin A in coffee Marta H. Taniwaki Mycobiota, mycotoxigenic fungi, and citrinin production in black olives Dilek Heperkan, Burçak E. Meriç, Gülçin Sismanoglu, Gözde Dalkiliç, and Funda K. Güler Byssochlamys: significance of heat resistance and mycotoxin production Jos Houbraken, Robert A. Samson and Jens C. Frisvad Effect of water activity and temperature on production of aflatoxin and cyclopiazonic acid by Aspergillus flavus in peanuts Graciela Vaamonde, Andrea Patriarca and Virginia E. Fernández Pinto

ECOPHYSIOLOGY OF FUMONISIN PRODUCERS IN FUSARIUM SECTION LISEOLA Vicente Sanchis, Sonia Marín, Naresh Magan and Antonio J. Ramos*

1.

INTRODUCTION

Fumonisins were first described as being produced by Fusarium Section Liseola species (Gelderblom et al., 1988; Marasas et al., 1988), and subsequently a number of toxicological studies have demonstrated their role in animal health problems caused by consumption of contaminated feeds. A relationship has been postulated between frequent ingestion of fumonisin containing maize and incidence of esophageal cancer by humans in certain areas of the world. Thus fumonisins have been classified as possible human carcinogens (Group 2B), by IARC (1993). High levels of fumonisins have been reported in maize in Africa, Asia and South America (Chu and Li, 1994; Doko et al., 1996; Kedera et al., 1999; Ono et al., 1999; MedinaMartinez and Martinez, 2000), sometimes co-occurring with other mycotoxins. Surveys have shown that much of the maize intended for human consumption is contaminated with fumonisins to some extent (Pittet et al., 1992; Sanchis et al., 1994), and these mycotoxins may contaminate a wide range of corn-based foods in our diet (Weidenboerner, 2001). * V. Sanchis ([email protected]), S. Marín and A. J. Ramos, Food Technology Dept, Lleida University, 25198 Lleida, Spain; N. Magan, Cranfield Biotechnology Centre, Cranfield University, Silsoe, England.

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IMPORTANCE OF THE ECOPHYSIOLOGICAL STUDIES

The importance of the widespread contamination of foods and feeds by fumonisins has led to a proliferation of studies aimed at understanding the ecophysiology of the Fusarium spp. involved, particularly F. verticillioides and F. proliferatum, and the delineation of environmental conditions that allow production of fumonisins. An understanding of the influence of biotic and abiotic factors on germination, growth and fumonisin production by these species is important in managing the problem of fumonisin contamination in the food supply. This study focuses on the impact of different abiotic factors including substrate, water activity (aw), temperature and preservatives, and biotic factors such as the natural mycoflora present in the grain.

3.

INFLUENCE OF ABIOTIC FACTORS ON FUNGAL DEVELOPMENT

3.1.

Substrate

Fusarium Section Liseola species are much more commonly found in maize than other grains, such as wheat and barley. Studies carried out by Marín et al. (1999a) have shown that even though Fusarium verticillioides and F. proliferatum are able to grow on a wide variety of substrates, including wheat, barley and maize, high fumonisin biosynthesis only occurs in maize. The assayed isolates were able to grow in the different cereal grains under similar conditions of temperature and aw, but negligible amounts of fumonisin B1 (FB1) were detected. Although fumonisins are found mainly in corn and corn-based foods and feeds, there are a few reports of fumonisins from other substrates such as ‘black oats’ animal feed from Brazil (Sydenham et al., 1992), New Zealand forage grass (Scott, 1993), Indian sorghum (Shetty and Bhat, 1997), rice (Abbas et al., 1998), asparagus (Logrieco et al., 1998), beer (Torres et al., 1998), wheat/barley/soybeans (Castella et al., 1999), and tea (Martins et al., 2001).

3.2.

Water Activity and Temperature

Our results show that germination of F. verticillioides and F. proliferatum is possible between 5-37°C at aw values above 0.88, but the

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range for growth is slightly narrower (7-37°C above 0.90 aw) (Figure 1). The lag phases were shorter at 25-30°C and 0.94-0.98 aw, and they increased to 10-500 h at marginal temperatures (5-10°C). There were some differences between strains. FB1 production in grain was observed between 10-37°C but only at aw of 0.93 and above. The optimum conditions for fumonisin production by F. verticillioides and F. proliferatum were 15-30°C at 0.97 aw (Marín et al., 1996; Marín et al., 1999b). These two environmental conditions (temperature and moisture availability) are the main factors which control fumonisin production in grain. The authors have developed detailed two-dimensional profiles of conditions that allow the production of FB1 (Marín et al., 1999b) (Figure 2).

3.3.

Preservatives

Grain preservatives based on propionates have shown some activity in controlling the growth of the Fusarium spp., and FB1 production. Growth rates decreased as preservative concentration increased, regardless of aw, while fumonisin production decreased only when aw was 0.93 or lower. In general, only a concentration ≥ 0.07% propionate was effective. In the presence of low propionate concentrations (0.03%), FB1 production was sometimes stimulated, possibly due to assimilation of these compounds by the moulds. The inhibitory effect of the preservatives is significantly affected by the water activity and temperature of the grain (Marín et al., 1999c).

Growth rate (mm d-1)

6 5 4 3 2 1 0

10 20 30 Temperature (°C)

40

Figure 1. Effect of temperature and water activity on growth rate of F. verticillioides. aw : 0.98 (●), 0.96 (▲), 0.94 (■), 0.92 (●), 0.90 (▲), 0.88 ( ■)

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Water activity

0.98 0.97 0.96 1000

0.95

100 50 10 5 1

0.94 0.93 0.92 5

10

15 20 25 30 Temperature (°C)

35

40

Figure 2. Water activity/temperature profiles for FB1 accumulation by F. verticillioides after 28 days of incubation on irradiated maize (from Marín et al., 1999b).

Essential oils from plants have been also tested for their antimycotoxigenic activity and found to inhibit FB1 accumulation in maize under moist conditions. Resveratrol, a compound known for its antioxidant properties, decreased FB1 accumulation in maize at a concentration as low as 23 ppm (Fanelli et al., 2003).

4.

INFLUENCE OF BIOTIC FACTORS ON FUNGAL DEVELOPMENT

In general, the presence of other fungal species in mixed cultures inhibited the growth of F. verticillioides and F. proliferatum. Aspergillus niger and A. flavus were particularly effective in decreasing the competitiveness of the Fusaria (Marín et al., 1998). However, their effectiveness depended on abiotic factors. In general, the Fusarium spp. competed better at higher aw levels (0.98), and temperatures close to 15°C. Interestingly, FB1 production was stimulated by certain species at high water availabilities (0.98 aw), mainly when competing with A. niger for occupation of the same niche (Table 1). Fumonisins are secondary metabolites: if Fusarium is an endophyte, fumonisin production may be more important for retaining its niche than for occupying it. However, if contamination occurs from the air

+ A. niger + A. ochraceus + A. flavus + P. implicatum F. proliferatum + A. niger + A. ochraceus + A. flavus + P. implicatum

0.2±0.1 0.6±0.0 12.2±6.5 0.1±1.0 17.2±4.8

0.4±0.1 1.1±0.1 15.8±18.6 0.2±0.1 34.0±8.8

137.9±1.4 84.3±1.8 12.8±6.0 3.8±1.6 22.8±6.7

0.5±0.0 0.3±0.2 0.2±0.1 11.5±15.6 5.6±1.0

5.1±0.3 47.3±2.9 0.2±0.0 0.7±0.1 3.6±4.7

360.9±9.2 40.2±0.6 10.4±7.1 48.0±11.9 0.7±0.5

19.1±26.4 0.1±0.0 6.8±2.7 3.6±4.5

33.5±6.3 1.7±0.1 105.9±1.6 284.3±401.4

1084.4±33.5 48.8±0.5 602.7±8.1 234.4±308.6

0.2±0.1 0.9±0.0 6.8±8.3 3.3±3.0

3.0±1.4 2.9±0.1 9.8±3.6 21.3±28.5

0.8±0.1 0.2±0.0 64.1±32.9 11.6±6.6

Ecophysiology of Fumonisin Producers in Fusarium Section Liseola

Table 1. Influence of water activity and temperature on production of fumonisin B1 (ppm) by Fusarium spp. on maize grain in the presence of competing mycoflora after a 4-weeks incubation period Temperature (°C) 15°C 25°C Water activity 0.98 0.95 0.95 0.93 0.98 0.93 F. verticillioides 54.1±10.7 7.1±1.3 29.3±0.4 0.8±0.1 5.5±5.9 4.8±5.2

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or leaves, establishment may be assisted by fumonisin production. However, field data do not support the hypothesis that F. verticillioides gains a competitive advantage via FB1 (Reid et al., 1999).

5.

CONCLUSION

Most fumonisin is produced in maize pre-harvest, but fumonisin control in maize post-harvest can be achieved by effective control of the moisture content (aw). Temperature control and periodic aeration, along with the natural microflora may act as additional controls.

6.

ACKNOWLEDGMENTS

The authors are grateful to the Spanish Government (CICYT, Comision Interministerial de Ciencia y Tecnologia, grant ALI98 0509C04-01) and the European Union (QLRT-1999-996) for the financial support.

7.

REFERENCES

Abbas, H. K, Cartwright, R. D., Shier, W. T., Abouzied, M. M., Bird, C. B., Tice, L. G., Ross, P. F., Sciumbato, G. L., and Meredith, F. I., 1998, Natural occurrence of fumonisins in rice with sheath rot disease, Plant Dis. 82:22-25. Castella, G., Bragulat, M. R., and Cabanes, F. J., 1999, Surveillance of fumonisins in maize-based feeds and cereals from Spain, J. Agric. Food Chem. 47:4707-4710. Chu, F. S., and Li, G. Y., 1994, Simultaneous occurrence of fumonisin B1 and other mycotoxins in moldy corn collected from the People’s Republic of China in regions with high incidences of esophageal cancer, Appl. Environ. Microbiol. 60:847-852. Doko, M. B., Canet, C., Brown, N., Sydenham, E. W., Mpuchane, S., and Siame, B. A., 1996, Natural co-occurrence of fumonisins and zearalenone in cereals and cereal-based foods from eastern and southern Africa, J. Agric. Food Chem. 44:3240-3243. Fanelli, C, Taddei, F., Trionfetti, P., Jestoi, M., Ricelli, A., Visconti, A., and Fabbri, A., 2003, Use of resveratrol and BHA to control fungal growth and mycotoxin production in wheat and maize seeds, Aspects Appl. Biol. 68:63-71. Gelderblom, W. C. A., Jaskiewicz, K.; Marasas, W. F. O., Thiel, P. G., Hora, R. M., Vleggaar, R., and Kriek, N. P. J., 1988, Fumonisins -novel mycotoxins with cancerpromoting activity produced by Fusarium moniliforme, Appl. Environ. Microbiol. 54:1806-1811.

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Kedera, C. J., Plattner, R. D., and Desjardins, A. E., 1999, Incidence of Fusarium spp. and levels of fumonisin B1 in maize in western Kenya, Appl. Environ. Microbiol. 65:41-44. IARC, 1993, IARC monographs on the evaluation of carcinogenic risks of chemicals to humans; some naturally occurring substances: food items and constituents, heterocyclic aromatic amines and mycotoxins. Ochratoxin A, International Agency for Research on Cancer, Lyon, pp. 26-32. Logrieco, A., Doko, M. B., Moretti, A., Frisullo, S., and Visconti, A., 1998, Occurrence of fumonisin B1 and B2 in Fusarium proliferatum infected asparagus plants, J. Agric. Food Chem. 46:5201-5204. Marasas, W. F. O., Jaskiewicz, K., Venter, F. S., and van Schalkwyk, D. J., 1988, Fusarium moniliforme contamination of maize in oesophageal cancer areas in Transkei, Sth Afr. Med. J., 74:110-114. Marín, S., Sanchis, V., Teixido, A., Saenz, R., Ramos, A. J., Vinas, I., and Magan, N., 1996, Water and temperature relations and microconidial germination of Fusarium moniliforme and F. proliferatum from maize, Can. J. Microbiol. 42:1045-1050. Marín, S., Sanchis, V., Rull, F., Ramos, A. J., and Magan, N., 1998, Colonization of maize grain by Fusarium moniliforme and Fusarium proliferatum in the presence of competing fungi and their impact on fumonisin production, J. Food Prot. 61: 1489-1496. Marín, S., Magan, N., Serra, J., Ramos, A. J., Canela, R., and Sanchis V., 1999a, Fumonisin B1 production and growth of Fusarium moniliforme and Fusarium proliferatum on maize, wheat, and barley grain, J. Food Sci. 64:921-924. Marín, S., Magan, N., Belli, N., Ramos, A. J., Canela, R., and Sanchis V., 1999b, Two dimensional profiles of fumonisin B1 production by Fusarium moniliforme and Fusarium proliferatum in relation to environmental factors and potential for modelling toxin formation in maize grain, Int. J. Food Microbiol. 51:159-167. Marín, S., Sanchis, V., Sanz, D., Castel, I., Ramos, A. J., Canela, R., and Magan, N., 1999c, Control of growth and fumonisin B1 production by Fusarium verticillioides and Fusarium proliferatum isolates in moist maize with propionate preservatives, Food Addit. Contam. 16:555-563. Martins, M. L., Martins, H. M., and Bernardo, F., 2001, Fumonisins B1 and B2 in black tea and medicinal plants, J. Food Prot. 64:1268-1270. Medina-Martinez, M. S., and Martinez, A. J., 2000, Mold occurrence and aflatoxin B1 and fumonisin B1 determination in corn samples in Venezuela, J. Agric. Food Chem. 48:2833-2836. Ono, E. Y. S., Sugiura, Y., Homechin, M., Kamogae, M., Vizzoni, E., Ueno, Y., and Hirooka, E.Y., 1999, Effect of climatic conditions on natural mycoflora and fumonisins in freshly harvested corn of the state of Parana, Brazil, Mycopathologia, 147:139-148. Pittet, A., Parisod, V., and Schellenberg, M., 1992, Occurrence of fumonisins B1 and B2 in corn-based products from the Swiss market, J. Agric. Food Chem. 40:1445-1453. Reid, L. M., Nicol, R. W., Ouellet, T., Savard, M., Miller, J. D., Young, J. C., Stewart, D. W., and Schaafsma, A. W., 1999, Interaction of Fusarium graminearum and F. moniliforme in maize ears: disease progress, fungal biomass, and mycotoxin accumulation, Phytopathology, 89:1028-1037. Sanchis, V., Abadias, M., Oncins, L., Sala, N., Vinas, I., and Canela, R., 1994, Occurrence of fumonisins B1 and B2 in corn-based products from the Spanish market, Appl. Environ. Microbiol. 60:2147-2148.

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Scott, P. M., 1993, Fumonisins, Int. J. Food Microbiol. 18:257-270. Shetty, P. H., and Bhat, R. V., 1997, Natural occurrence of fumonisin B1 and its co-occurrence with aflatoxin B1 in Indian sorghum, maize and poultry feeds, J. Agric. Food Chem. 45:2170-2173. Sydenham, E. W., Marasas, W. F. O., Shephard, G. S., Thiel, P. G., and Hirooka, E. Y., 1992, Fumonisin concentrations in Brazilian feeds associated with field outbreaks of confirmed and suspected animal mycotoxicoses, J. Agric. Food Chem. 40: 994-997. Torres, M. R., Sanchis, V., and Ramos, A. J., 1998, Occurrence of fumonisins in Spanish beers analyzed by an enzyme-linked immunosorbent assay method, Int. J. Food Microbiol. 39:139-143. Weidenboerner, M., 2001, Foods and fumonisins, Eur. Food Res. Technol. 212: 262-273.

ECOPHYSIOLOGY OF FUSARIUM CULMORUM AND MYCOTOXIN PRODUCTION Naresh Magan, Russell Hope and David Aldred *

1.

INTRODUCTION

Fusarium ear blight is a cereal disease responsible for significant reduction in yield and quality of wheat grain throughout the world. In addition to degradation in grain quality, Fusarium species produce an array of mycotoxins which may contaminate the grain. This mycotoxin production occurs preharvest and during the early stages of drying (Botallico and Perrone, 2002; Magan et al., 2002). F. culmorum is the most common cause of Fusarium ear blight in the United Kingdom and some other countries and can produce trichothecenes including deoxynivalenol (DON) and nivalenol (NIV). DON and NIV are harmful to both animals and humans, causing a wide range of symptoms of varying severity, including immunosuppression. Germination of macroconidia of F. culmorum can occur over a wide range of temperatures (5-35°C) with a minimum aw near 0.86 at 20-25°C based on an incubation period of about 40 days (Magan and Lacey, 1984a). Longer term incubations on other media have suggested limits for germination of about 0.85 aw (Snow, 1949). The ecological strategies used by F. culmorum to occupy and dominate in the grain niche are not understood. Fungi can have combative (C-selected), stress (S-selected) or ruderal (R-selected) strategies or * Applied Mycology Group, Biotechnology Centre, Cranfield University, Silsoe, Bedford MK45 4DT, U.K. Correspondence to [email protected]

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merged secondary strategies (C-R, S-R, C-S, C-S-R; Cooke and Whipps, 1993). Thus primary resource capture of nutritionally rich food matrices such as grain by F. culmorum may depend on germination and growth rate, enzyme production, sporulation and the capacity for producing secondary metabolites to enable effective competition with other fungi. Attempts to control F. culmorum and other Fusarium species have relied on the application of fungicides preharvest coupled with effective storage regimens. However, the timing and application of these control measures are critical. Some fungicides are ineffective against Fusarium ear blight and may in some cases result in a stimulation of mycotoxin production, particularly under suboptimal fungal growth conditions and low fungicide doses (D’Mello et al., 1999; Jennings et al., 2000; Magan et al., 2002). It has been shown that moisture conditions at anthesis are crucial in determining infection and mycotoxin production by F. culmorum on wheat during ripening. Few studies have been carried out to determine the effect that key environmental factors such as water activity (aw), temperature and time have on fungal growth and mycotoxin production. Some studies have identified the aw range for germination and growth of F. culmorum and other Fusarium species (Sung and Cook, 1981; Magan and Lacey 1984a; Magan and Lacey 1984b). The combined effect of aw and temperature has been found to be significant for growth and fumonisin production by Fusarium verticillioides and F. proliferatum which infect maize (Marin et al. 1999). Knowledge of the threshold limits for growth and mycotoxin production are very important in controlling the entry of mycotoxins into the food chain. The objective of this study was to examine in detail the impact of water availability, temperature and time on growth and DON and NIV production by an isolate of F. culmorum on a medium based on wheat grains. Production of enzymes that may assist F. culmorum to dominate in the grain niche was also examined.

2.

MATERIALS AND METHODS

2.1.

Fungal Isolates and Media

A representative strain of Fusarium culmorum (98WW4.5FC, Rothamsted Research Culture Collection, Harpenden, Herts, UK) was chosen from a range examined previously and isolated from UK

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wheat grain, with a known history of mycotoxin production (Lacey et al., 1999; Magan et al., 2002). The strain produced quantities of mycotoxins comparable with other strains from Europe. A 2% milled wheat grain agar (Agar No. 3, Oxoid, Basingstoke, UK) was used as the basic medium. The aw was adjusted with glycerol in the range 0.995-0.850 as described by Dallyn (1978). The equivalent moisture contents of the aw treatments of 0.995, 0.98, 0.95, 0.90 and 0.85 were 30%, 26%, 22%, 19% and 17.5% for wheat grain. Glycerol was used because of its inherent aw stability over the temperature range 10-40°C. Media were sterilised by autoclaving for 15 min at 120°C. Media were cooled to 50°C before pouring into 90 mm Petri plates. The aw of media was confirmed using an Aqualab instrument (Decagon Inc., Washington State, USA). In all cases the aw levels were checked at both incubation temperatures (15° and 25°C) and were within 0.003 of the desired levels. For studies in whole grain, winter wheat was gamma irradiated at 12kGy to remove contaminant microorganisms, but conserve germinative capacity. No mycotoxins were found in the grain lot used. Varying amounts of water were added to the grain and an adsorption curve prepared to facilitate accurate modifications of the aw of the grain comparable with the media-based studies. Grain was placed in sterile flasks and inoculated with appropriate volumes of sterile water to obtain the necessary treatments. The flasks were sealed and left for 24-36 h to equilibrate at 4°C. The grain was then decanted carefully into 90 mm Petri dishes to obtain a monolayer of wheat grain. In all cases the aw levels were checked at both temperatures as described above and were within 0.003 of the desired levels.

2.2.

Inoculation and Growth Measurements

For both agar and grain based studies, replicates of each aw treatment were inoculated centrally with a 5µl drop of a 105 cfu/ml F. culmorum macroconidial suspension obtained from a 7 d colony grown on 2% milled wheat agar. Conidia were obtained by flooding the culture with 5 ml sterile distilled water containing 0.5% Tween 80 and agitating the colony surface with a sterile glass rod. Replicates of the same treatment were enclosed in polyethylene chambers together with 500 ml of a glycerol/water solution of the same aw, closed and incubated at 15° or 25°C for up to 40 days. Growth measurements were taken throughout the incubation period, by taking two diametric measurements of the colonies at right angles. Colonisation rates were determined subsequently by linear regression of the radial extension

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rates. Three replicates per treatment were removed after 10, 20, 30 or 40 d and analysed for DON and NIV (agar media) and for DON (wheat grain). The experiments were repeated once.

2.3.

Mycotoxin Extraction and Analyses

Mycotoxin extraction was adapted from Cooney et al. (2001). The entire agar and mycelial culture or grain sample from each replicate sample was placed in acetonitrile/methanol (14:1; 40 ml) and shaken for 12 h. Aliquots (2 ml) were taken for DON/NIV analysis and passed through a cleanup cartridge comprising a 2 ml syringe (Fisher Ltd.) packed with a disc of filter paper (No. 1 Whatman International Ltd.), a 5 ml luger of glass wool and 300 mg of alumina/activated carbon (20:1). The sample was allowed to gravity feed through the cartridge and residues in the cartridge washed out with acetonitrile/methanol/water (80:5:15; 500 µl). The combined eluate was evaporated (compressed air, 50°C) and then resuspended in methanol/water (5:95; 500 µl). Quantification of DON/NIV was accomplished by HPLC, using a Luna™ C18 reverse phase column (100 mm × 4.6 mm i.d.) (Phenomenex, Macclesfield, U.K.). Separation was achieved using an isocratic mobile phase of methanol/water (12:88) at an elution rate of 1.5 ml/min. Eluates were detected using a UV detector set at 220 nm with an attenuation of 0.01 AUFS. The retention times for NIV and DON were 3.4 and 7.5 min respectively. External standards were used for quantification (Sigma-Aldrich, Poole, Dorset, U.K.). The limit for quantification was 5 ng/g for DON and 2.5 ng/g for NIV.

2.4.

Hydrolytic Enzyme Profiles in Grain

For enzyme extraction subsamples of grain (2 g) were placed in 4 ml potassium phosphate extraction buffer (10 mM; pH 7.2). The bottles were shaken on a wrist action shaker for 1 h at 4°C. Washings were decanted into plastic Eppendorf tubes (1.5 ml) and centrifuged in a bench microcentrifuge for 15 min. The supernatant was decanted and stored in aliquots at −20°C for total and specific enzyme activity determinations. The total activity of seven hydrolytic enzyme activities was assayed using ρ-nitrophenyl substrates (Sigma Chemical Co., UK). Enzyme extract (40 µl), substrate

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solution (40 µl) and the appropriate buffer (20 µl) were pipetted into the wells of the microtitre plate and incubated at 37°C for 1 h along with the appropriate controls. The reaction was stopped by the addition of 5 µl 1M sodium carbonate solution and left for 3 min. The enzyme activity was measured, using a MRX multiscan plate reader (Dynex Technologies Ltd., Billinghurst, UK), by the increase in optical density at 405 nm caused by the liberation of ρ-nitrophenol by enzymatic hydrolysis of the substrate. Enzyme activity was calculated from a calibration curve of absorbance at 405 nm vs ρ-nitrophenol concentration and expressed as µmol ρ-nitrophenol released/min. For specific activity determinations the protein concentration was obtained using a Bicinchoninic acid protein assay kit (Sigma-Aldrich Ltd, Poole, Dorset, UK). This kit consisted of bicinchoninic acid solution, copper (II) sulphate pentahydrate 4% solution and albumin standard (containing bovine serum albumin (BSA) at a concentration of 1.0 mg/ml). Protein reduces alkaline Cu (II) to Cu (I), which forms a purple complex with bicinchoninic acid (a highly specific chromogenic reagent). The resultant absorbance at 550 nm is directly proportional to the protein concentration. The working reagent was obtained by the addition of 1 part copper (II) sulphate solution to 50 parts bicinchoninic acid solution. The reagent is stable for one day provided it is stored in a closed container at room temperature. Aliquots (10 µl) of each standard or enzyme extracts were placed in the appropriate microtitre plate wells. Potassium phosphate extraction buffer 10 mM pH 7.2 (10 µl) was pipetted into the blank wells. The working reagent (200 µl) was added to each well, shaken and plates incubated at 37°C for 30 min. The plates were allowed to cool to room temperature before measuring the absorbance at 550 nm using a MRX multiscan plate reader. The protein concentrations in the enzyme extracts were obtained from the calibration curve of absorbance at 550 nm against BSA concentration. These values were used to calculate the specific activity of the enzymes in nmol ρ-nitrophenol released per min per µg protein.

2.5.

Statistical Analyses

The data were analysed using ANOVA (SigmaStat, SPSS Inc.), with significance values of 30% M.C.

0.99

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5.0

0.97

3.0

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0.93 21-22%

Water activity/Moisture content

0.91 0.89

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(a) Growth rate (mm day-1) 10

0.99 1.0 0.97

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0.1 21-22%

0.93

0.01

0.91 0.89 0.87 18-19%

(b) Deoxynivalenol (ppm) 0.85 0

10

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30

40

Temperature (⬚C)

Figure 5. Comparison of profiles and limits for (a) germination ( ) and growth (mm/day) and (b) DON (mg/kg) production by Fusarium culmorum on wheat grain (compiled from Magan and Lacey, 1984a,b; Magan, 1988; Hope, 2003; Hope and Magan, 2003).

resources over a range of aw and temperature conditions. Previous studies have demonstrated that F. culmorum produces a significant amount of cellulases over a range of aw levels (Magan and Lynch, 1986). These hydrolytic enzymes may, when combined with secondary

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100

µmol 4-nitrophenol min−1 µg−1 grain

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60

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20

0 (a) 12

nmol 4-nitrophenol min−1 µg−1 protein

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(b)

β-D-fucosidase

α-Dgalactosidase

β-Dglucosidase

α-Dmanno sidase

β-D- N-acetyl-α- N-acetyl-βxylo- D-Glucos- D-Glucossidase aminidase aminidase

Figure 6. Production of total (a) and specific (b) activity of seven different enzymes by F. culmorum on irradiated wheat grain incubated for 14 days at 0.99 aw and 25°C (Hope, 2003).

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metabolite production and tolerance to intermediate moisture conditions facilitate competitiveness in food raw materials. Studies of F. verticillioides and F. proliferatum have also shown that production of some hydrolytic enzymes by these species during colonisation of maize may be used as an indicator and diagnosis of early infection by these species (Marin et al., 1998). It is of interest that F. culmorum appears less competitive than F. graminearum when the two species interact in vitro on agar media or on wheat grain (Magan et al., 2003), as both occupy similar ecological niches.

4.

CONCLUSIONS

Growth, competitiveness, dominance and mycotoxin production in food matrices are influenced by complex interactions between the environment, the prevailing fungal community, and external factors. The role of mycotoxins is still unclear, but they may enable fungi to occupy a particular niche, or assist in excluding other competitors from the same niche.

6.

REFERENCES

Bottalico, A., and Giancarlo, P., 2002, Toxigenic Fusarium species and mycotoxins associated with head blight in small-grain cereals in Europe, Eur. J. Plant Pathol. 108:611-624. Cooke, R. C., and Whipps, J. M., 1993. Ecophysiology of Fungi, Blackwell Scientific Publications, Oxford UK. 337 pp. Cooney, J. M., Lauren, D. R., and di Menna, M. E., 2001, Impact of competitive fungi on trichothecene production by Fusarium graminearum, J. Agric. Food Chem. 49:522-526. Dallyn, H., 1978, Effect of substrate, water activity on the growth of certain xerophilic fungi, PhD thesis, Polytechnic of the South Bank London, Council for National Academic Awards. D’Mello, J. P. F., Placinta, C. M., and Macdonald, A. M. C., 1999, Fusarium mycotoxins: a review of global implications for animal health, welfare and productivity, Animal Food Sci. Technol. 80:183-205. Jennings, P., Turner J. A., and Nicholson, P., 2000, Overview of Fusarium ear blight in the UK -effect of fungicide treatment on disease control and mycotoxin production, The British Crop Protection Council. Pests and Diseases 2000 2:707-712. Hope, R., 2003, Ecology and control of Fusarium species and mycotoxins in wheat grain, PhD thesis, Institute of BioScience and Technology, Cranfield University, Silsoe, Bedford, U. K.

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Hope, R. and Magan, N., 2003, Two-dimensional environmental profiles of growth, deoxynivalenol and nivalenol production by Fusarium culmorum on a wheat-based substrate, Lett. Appl. Microbiol. 37:70-74. Lacey J., Bateman G. L., and Mirocha C. J., 1999, Effects of infection time and moisture on development of ear blight and deoxynivalenol production by Fusarium spp. in wheat, Annals Appl. Biol. 134:277-283. Magan, N., 1988, Effect of water potential and temperature on spore germination and germ tube growth in vitro and on straw leaf sheaths, Trans. Br. Mycol. Soc. 90: 97-107. Magan, N., and Lacey, J., 1984a, The effect of temperature and pH on the water relations of field and storage fungi, Trans. Br. Mycol. Soc. 82:71-81. Magan, N., and Lacey, J., 1984b, Water relations of some Fusarium species from infected wheat ears and grain, Trans. Br. Mycol. Soc. 83:281-285. Magan, N., and Lynch, J.M., 1986, Water potential, growth and cellulolysis of soil fungi involved in decomposition of crop residues, J. Gen. Microbiol. 132:11811187. Magan, N., Hope, R., Colleate, A., and Baxter, E. S., 2002, Relationship between growth and mycotoxin production by Fusarium species, biocides and environment, Eur. J. Plant Pathol. 108:685-690. Magan, N., Hope, R., Cairns, V., and Aldred, D., 2003, Post-harvest fungal ecology: impact of fungal growth and mycotoxin accumulation in stored grain, Eur. J. Plant Pathol. 109:723-730. Marin, S., Sanchis, V., and Magan, N., 1998, Effect of water activity on hydrolytic enzyme production by F. moniliforme and F. proliferatum during early stages of growth on maize, Int. J. Food Microbiol. 42:1-10 Marin, S., Sanchis, V., Ramos, A. J., and Magan, N., 1999, Two-dimensional profiles of fumonisin B1 production by Fusarium moniliforme and F. proliferatum in relation to environmental factors and potential for modelling toxin formation in maize grain, Int. J. Food Microbiol. 51:159-167. Northolt, M. D., van Egmond, H. P., and Paulsch, W. E., 1979, Ochratoxin A production by some fungal species in relation to water activity and temperature, J. Food Prot. 42:485-490. Northolt, M. D., Verhulsdonk, C. A. H., Soentoro, P. S. S. and Paulsch, W. E., 1976, Effect of water activity and temperature on aflatoxin production by Aspergillus parasiticus, J. Milk Food Technol. 39:170-174. Ramirez, M. L., Chulze, S. N., and Magan, N., 2004, Impact of environmental factors and fungicides on growth and deoxinivalenol production by Fusarium graminearum isolates from Argentinian wheat, Crop Prot. 23:117-125. Snow, D., 1949, Germination of mould spores at controlled humidities, Annals Appl. Biol. 36:1-13. Sung, J. M., and Cook, R. J., 1981, Effect of water potential on reproduction and spore germination of Fusarium roseum “graminearum”, “culmorum” and “avenaceum,” Phytopathology 71:499-504.

FOOD-BORNE FUNGI IN FRUIT AND CEREALS AND THEIR PRODUCTION OF MYCOTOXINS Birgitte Andersen and Ulf Thrane*

1.

INTRODUCTION

The growth of filamentous fungi in foods and food products results in waste and is costly as well as sometimes hazardous. Many different fungal species can spoil food products or produce mycotoxins or both. As each fungal species produces its own specific, limited number of metabolites and is associated with particular types of food products, the number of mycotoxins potentially present in a particular product is limited (Filtenborg et al., 1996). If physical changes occur in a product, changes in the association of fungal species found in the product will also occur. With current understanding it is possible to predict which fungi and mycotoxins a given product may contain, when the type of food product and the history of production and storage are known. In Europe, fruit has received minor attention in relation to fungal spoilage, whereas fungal spoilage of cereals has been studied extensively, but often with the focus on only one or two fungal genera. Apple juice is one of the few commodities that has caught the attention of the European authorities and regulation of patulin will be in force by the end of 2003 in Denmark (EC, 2004).

* Center for Microbial Biotechnology, BioCentrum-DTU, Technical University of Denmark, DK-2800 Kgs. Lyngby, Denmark. Correspondence to: ba@biocentrum. dtu.dk

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Knowledge of the composition and succession of the mycobiota in cereal grains and fruit during maturation, harvest and storage is an important step towards the prediction of possible mycotoxin contamination. Some major spoilage genera on stored apples and cherries, e.g. Botrytis, Cladosporium and Rhizopus are not known to produce significant mycotoxins, while others including Alternaria, Aspergillus, Fusarium and Penicillium include species capable of producing a wide range of mycotoxins (Pitt and Hocking, 1997). The production of mycotoxins is often species specific (Frisvad et al., 1998), so accurate identification of fungi to species is of major importance. Historically, identifications have not always been accurate, and incorrect identifications have resulted in confusion and misinterpretations (Marasas et al., 1984; Thrane, 2001; Andersen et al., 2004). In the case of Alternaria, where many taxa are still undescribed (Simmons and Roberts, 1993; Andersen et al., 2002), identification is only possible to a species-group level for many isolates. Before a contaminated sample is analysed for mycotoxins, it is important to know which mycotoxins are likely to be present. Metabolite profiles from known species grown in pure culture can provide valuable information about the mycotoxins that may be found in cereals and fruit and their products, once the fungi normally associated with those products are known. During the last 15 years our group has analysed numerous cereal and fruit samples and recorded the fungal species found. Analysis of that large amount of data has shown that similar fungal species occur on the same product types year after year. The purpose of this paper is to present a list of the fungal species found on apples, cherries, barley and wheat from the Northern temperate zone together with a list of mycotoxins known to be produced by these fungi.

2.

MATERIALS AND METHODS

2.1.

Media

Dichloran Rose Bengal Yeast Sucrose agar (DRYES; Frisvad, 1983) and V8 juice agar (V8; Simmons, 1992) were used for fungal analyses of fruit, while Czapek Dox Iprodione Dichloran agar (CZID; Abildgren et al., 1987), Dichloran 18% Glycerol agar (DG18; Hocking and Pitt, 1980), DRYES, and V8 were used for analysis of cereals. CZID plates were incubated in alternating light/dark cycle

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consisting of 12 hours of black fluorescent and cool white daylight and 12 hours darkness at 20-23°C, while DG18 and DRYES plates were incubated at 25°C in darkness and V8 plates in alternating cool white daylight (8 hours light/16 hours darkness) at 20-23°C. Alternaria and other dematiaceous hyphomycetes were enumerated on DRYES and/or V8. Fusarium species were enumerated on CZID and/or V8, while Eurotium, Aspergillus and Penicillium species and other hyaline fungi were enumerated on DG18 and/or DRYES. A wide range of media were used for fungal identification. For Alternaria species and other black fungi, DRYES, Potato Carrot Agar (PCA; Simmons, 1992) and V8 were used; for Eurotium species, Malt Extract Agar (MEA, Pitt and Hocking, 1997) and Czapek Dox agar (CZ; Samson et al., 2004) were used; for Fusarium species, Potato Dextrose agar (PDA; Samson et al., 2004) Yeast Extract Sucrose agar (YES; Samson et al., 2004) and Synthetischer nährstoffarmer agar (SNA: Nirenberg, 1976) were used. For Penicillium, Czapek Yeast extract Agar (CYA; Pitt and Hocking, 1997), MEA, YES and Creatine Sucrose agar (CREA; Samson et al., 2004) were used.

2.2.

Fruit

Apple flowers with petals removed, apple peel and apples with fungal lesions were plated directly onto DRYES and V8. Sound apples were surfaced disinfected by shaking in freshly prepared 0.4 % NaOCl for 2 minutes and then rinsing with sterile water. The cores of the surface disinfected apples were cut out with a sterile cork borer, cut into 5 pieces and plated on V8 and DRYES. Cherry flowers with petals and cherries with fungal lesions were plated directly onto DRYES and V8. Some cherries were surfaced disinfected as described above. The stem and calyx ends of the surface disinfected cherries were excised with a sterile scalpel and plated on V8 and DRYES. The plates were incubated as described above and after a week of incubation the fungal colonies were enumerated. Representative colonies were then isolated and identified to species level. The development in the mycobiota at genus level was followed on apples (variety Jonagored) and two varieties of cherries (Vicky and Van) during one growth season (2001 for cherries and 2002 for apples). The trees were sprayed for fungal diseases (grey mould, Botrytis spp.) several time before harvest. Flowers of both apples and cherries were examined for fungal growth before the first spray application. Ten trees were selected from each orchard and sampled (10 units) from each tree. The ten samples from each orchard were

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pooled (100 units per sample) and screened for the presence of Alternaria, Botrytis, Cladosporium, Fusarium and Penicillium species.

2.3.

Cereals

Barley and wheat samples were plated directly with and without surface disinfection. Kernels were surfaced disinfected by shaking in freshly prepared 0.4 % NaOCl for 2 minutes and then rinsing with sterile water. Each sample consisted of 500 grains, that were not surface disinfected. All samples were plates on DG18, DRYES, V8 and CZID. The plates were incubated as described above and after a week of incubation the fungal colonies were enumerated. Representative colonies were then isolated and identified to species level. They were screened for the presence of Alternaria, Bipolaris, Cladosporium, Eurotium/Aspergillus, Fusarium and Penicillium species. The development in the mycobiota at genus level was followed on wheat (variety Leguan) and barley (variety Ferment) during one growth season (2002). One batch of barley seed was treated with fungicide before planting, while a second batch, and the wheat seed, were not treated.

2.4.

Fungal Species Associated with Fruit and Cereals

In our laboratory, data on the occurrence of fungal contamination of fresh, stored, processed and mouldy samples of cereals and fruits have been collected, recorded and compiled for more than 15 years. Samples from the field, food factories and local supermarkets have been surveyed. A number of cultivars of apples (e.g. Cox’s Orange, Discovery, Gala, Jonagored), cherries (e.g. Bing, Van, Vicky), barley (e.g. Alexis, Chariot, Ferment, Krona) and wheat (e.g. Leguan) have been sampled and the resulting data analysed.

2.5.

Fungal Identification and Metabolite Profiling

Alternaria and Stemphylium isolates were transferred to DRYES and PCA, while other black fungi, including Cladosporium isolates, were transferred to DRYES and V8. Identifications of the black fungi were done according to Andersen et al. (2002), Simmons and Roberts (1993), Samson et al. (2004) and Simmons (1967; 1969; 1986). Metabolite profiling involved extracting nine plugs of 14 day old DRYES cultures in ethyl acetate (1 ml) with formic acid (1%; Andersen et al., 2002) in an ultrasonic bath for 60 min. The ethyl

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acetate was evaporated and the dried sample redissolved in methanol (500 µl). Samples were then filtered and analysed on a HP1100 HPLCDAD (Agilent, Germany) (Andersen et al., 2002). Fusarium isolates were transferred to SNA, PDA and YES and identified according to Gerlach and Nirenberg (1982), Nirenberg (1989), Burgess et al. (1994) and Samson et al. (2004). Metabolites were profiled by extracting nine plugs of 14 day old PDA and YES cultures, each in dichloromethane: ethyl acetate (2:1 vol/vol; 1 ml) with formic acid (1%) in an ultrasonic bath for 60 min. Then the organic phase was evaporated and the dried sample redissolved in methanol (500 µl). Samples were then filtered and analysed on a HP1100 HPLCDAD (Smedsgaard, 1997). Penicillium isolates were transferred to CYA, MEA and YES and identified according to Samson et al. (2004) and Samson and Frisvad (2004). Metabolites were profiled by extracting three plugs of 7 day old CYA and YES cultures, then treated as for Fusarium extracts.

2.6.

Metabolites from Mouldy Fruit and Cereals

Metabolites were extracted from the mouldy samples in the same way as for the pure fungal cultures. Each sample (100 g) was mixed with ethyl acetate (100 ml) containing formic acid (1%). The mixture was shaken regularly over a 2 hour period to extract the metabolites, then held overnight in a freezer. Extracts (14 ml) were decanted from the frozen water and sample matrix, evaporated to dryness, redissolved in methanol (500 µl) and analysed as before.

3.

RESULTS

3.1.

Fungal Development in Fruit

The dominant fungal genera found in flowers, immature and mature fruit are given in Table 1. In apples, Cladosporium and Alternaria species constituted the major infection in the flowers. Botrytis and Fusarium species were also found frequently, whereas Penicillium species were isolated only rarely. After the trees were sprayed with fungicide, Botrytis was eliminated and the number of Cladosporium colonies was halved. The spraying did not seem to effect the numbers of Alternaria, Fusarium or Penicillium colonies enumerated. The numbers of these three genera increased with time and were highest at harvest.

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Table 1. Fungal infection (%) in Danish apples and cherries during the growth seasons 2002 and 2001 respectively Sour Sweet Apples cherries cherries (Jonagored) (Vicky) (Van) Fl.a Imm.b Mat.c Fl. Imm. Mat. Fl. Imm. Mat. Alternaria 77 80 88 68 33 60 66 22 18 Botrytis 35 0 0 4 0 5 78 1 2 Cladosporium 81 73 40 26 0 31 37 5 21 Fusarium 20 53 64 2 0 7 0 1 6 Penicillium 5 13 20 0 0 0 0 1 0 a Fl. = flowers; b Imm. = immature; cMat. = mature

On the flowers of sweet cherries, Botrytis was the dominant genus, whereas it was isolated in very low numbers from the flowers of sour cherries (Table 1). In flowers of both cherry types, Alternaria and Cladosporium were found in high numbers and Fusarium was found in low numbers. After spraying, Botrytis was more or less eliminated in both immature and mature cherries. The number of Cladosporium colonies seen was greatly reduced in immature cherries after spraying, but the numbers rose again as the cherries matured. Numbers of Alternaria isolated were somewhat reduced in immature cherries after spraying, but the numbers rose again in sour cherries while it fell in sweet cherries as they matured. A Penicillium species was found in only one sample of immature sweet cherries. The dominant toxigenic fungi on apples were Alt. tenuissima species-group, followed by Alt. arborescens species-group, F. avenaceum, F. lateritium, P. crustosum and P. expansum. On cherries, the dominant species were Alt. arborescens species-group followed by Alt. tenuissima species-group, F. lateritium and P. expansum.

3.2.

Fungal Development in Cereals

The dominant genera found in seed, immature (harvested by hand) and mature (machine harvested) wheat and barley kernels are shown in Table 2. In untreated wheat seed, Penicillium constituted the major infection in the seed together with Alternaria, Eurotium and Aspergillus. The same composition of fungi was seen in untreated barley seed, except that Penicillium counts were less than 50% of those in treated seed. In barley seeds that had been treated with fungicides before sowing, only two out of 500 grains were found to be infected with fungi. The changes in mycobiota in immature kernels compared to the seed were most pronounced in barley, where the numbers of

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Table 2. Fungal infection (%) in Danish wheat and barley seeds and kernels without surface disinfection during the growth season 2002 Wheat Barley Barley (Leguan) (Ferment) (Ferment) Seeda Imm.b Mat.c Seeda Imm. Mat. Seedd Imm. Mat. Alternaria 42 42 67 40 73 64 0 88 58 Botrytis 0 0 9 5 43 37 0 22 31 Eurotium / 25 0 1 19 1 0 0 1 0 Aspergillus Cladosporium 0 20 2 0 94 0 1 12 18 Fusarium 0 82 90 0 59 83 0 78 98 Penicillium 98 89 98 41 7 98 1 0 98 a Seed without fungicide treatment; b Imm. = Immature; c Mat. = Mature; d Fungicide treated seed

Fusarium and Alternaria rose markedly and the storage fungi disappeared. The change was less dramatic in wheat. The differences in mycobiota from immature to mature kernels were minor and mostly the number of fungal infected kernels was stable or rose slightly. The number of Cladosporium found on immature and mature samples varied a great deal as high number of Fusarium and Penicillium colonies often obscured the smaller Cladosporium colonies. Surface disinfection reduced the numbers of Penicillium and Eurotium colonies by 80-90 %, and of Cladosporium and Fusarium by 40-50 %. Only 10-15% of Alternaria and Bipolaris could be removed, indicating that the grains had internal infections with these genera. Dominant fungi in common to both wheat and barley kernels were isolates of Alt. infectoria species-group, F. avenaceum, P. aurantiogriseum, P. cyclopium and P. polonicum. However, barley had higher infection rates with Bipolaris sorokiniana, P. hordei and P. verrucosum compared to wheat.

3.3.

Fungal Species Associated with Fruit and Cereals

The frequencies of occurrence of fungal species in several cultivars of apples, cherries, barley and wheat are given in Tables 3 and 4. In our laboratory, such data have been compiled for more than 15 years. The differences in the mycobiota between cultivars were in most cases small. Most often the same fungal species were found and the variation was quantitative only. As can been seen from Tables 3 and 4, only a limited number of fungal species are found in both fruit and cereals. Newly harvested, undamaged apples and cherries were not usually infected by fungi. When infection was present, fungi mostly belonged

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to the dematiaceous hyphomycetes (Table 3). The mycobiota of fresh apples consisted of mainly of Alt. tenuissima species-group, Botrytis and Cladosporium spp. In fresh cherries, Alt. arborescens speciesgroup and Stemphylium spp. constituted the mycobiota, along with Botrytis and Cladosporium spp. In storage, the mycobiota changed in both types of fruit. In cherries, after only a few weeks in storage, Botrytis spp., P. expansum and Zygomycetes dominated the mycobiota and they often spoiled the cherries. In stored apples the same fungal species were found after months of storage together with P. solitum. Fungi were rarely found in juice made from apples or cherries, but when fungal growth was detected, Byssochlamys spp. and P. expansum were found in pasteurised and untreated juices, respectively. In contrast to fruit, samples of newly harvested, sound cereal grains always had some fungal infections after surface disinfection. Colonies of Alt. infectoria species-group, Cladosporium spp., F. avenaceum and F. tricinctum were always isolated from fresh barley and wheat. Epicoccum nigrum, F. culmorum, F, equiseti and F. poae were often seen also. As in fruit, the mycobiota changed in cereals in storage in favour of Aspergillus and Penicillium species. In dry barley and wheat samples that had been stored for one year or more, Eurotium spp. and Table 3. Fungal occurrence (frequency) in apples and cherries from the north temperate zone: data accumulated over a 15 year period a Apples Cherries Fresh Stored Juice Fresh Stored Juice Alternaria arborescens + − − ++ + − sp.-grp. Alt. infectoria sp.-grp. (+) − − (+) − − Alt. tenuissima sp.-grp. ++ + − + − − Botrytis spp. ++ ++ − ++ ++ − Byssochlamys spp. − − + − − + Cladosporium spp. ++ + − ++ + − Fusarium avenaceum + + − − − − F. lateritium + (+) − + − − Monilia spp. (+) + − (+) + − Penicillium carneum (+) + − − − − P. crustosum + + − − − − P. expansum + ++ (+) + ++ (+) P. polonicum − (+) − − − − P. solitum (+) ++ − − − − Stemphylium spp. (+) − − ++ − − Zygomycetes − ++ − − ++ − a +++: Always present; ++: often present; +: sometimes present; (+): rarely present; −: never detected or found only once. Fresh indicates direct plated, surface disinfected sound samples; stored, direct plated disinfected sound or visibly mouldy samples

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P. aurantiogriseum always dominated the mycobiota. However, Alt. infectoria species-group could still be isolated from barley that had been stored for two years. Aspergillus candidus, Asp. flavus, P. cyclopium, P. hordei, P. melanoconidium, P. polonicum, P. verrucosum and P. viridicatum were also often found in stored barley and in lower numbers in stored wheat.

3.4.

Production of Toxic Metabolites in Pure Culture

Six genera out of the twelve listed in Tables 3 and 4 are regarded as being non-toxigenic, namely Botrytis, Cladosporium, Epicoccum, Table 4. Fungal occurrence (frequency) in barley and wheat from the north temperate zone: data accumulated over a 15 year period a Barley Wheat Fresh Stored Fresh Stored Alternaria arborescens sp.-grp. (+) − (+) − Alt. infectoria sp.-grp. +++ ++ +++ ++ Alt. tenuissima sp.-grp. + − + − Aspergillus candidus − ++ − ++ Asp. flavus − ++ − ++ Asp. niger − + − + Bipolaris sorokiniana + (+) (+) − Botrytis spp. (+) − (+) − Cladosporium spp. +++ (+) +++ (+) Epicoccum nigrum ++ − ++ − Eurotium spp. − +++ − +++ Fusarium avenaceum +++ + +++ + F. culmorum ++ (+) ++ (+) F. equiseti ++ − ++ − F. graminearum + − + − F. langsethiae (+) − − − F. lateritium (+) − − − F. poae ++ + ++ + F. sporotrichioides + − + − F. tricinctum +++ + +++ + Penicillium aurantiogriseum (+) ++ (+) ++ P. cyclopium (+) ++ (+) ++ P. freii (+) + (+) ++ P. hordei + ++ (+) + P. melanoconidium − ++ − ++ P. polonicum − ++ − ++ P. verrucosum + ++ + ++ P. viridicatum − ++ − ++ Stemphylium spp. (+) − (+) − a See footnote to Table 3

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Eurotium, Monilia and Stemphylium, together with the Zygomycetes (Pitt and Hocking, 1997). Two genera which include toxigenic species, Bipolaris and Byssochlamys, were found only relatively rarely in cereals and fruit, respectively. Alternaria, Aspergillus, Fusarium and Penicillium species were much more commonly isolated. These four genera include 25 toxigenic species found on a regular basis in either fruit or cereals. Only Alt. tenuissima species-group and F. avenaceum were regularly found in both fresh fruit and fresh cereals. In Table 5 the mycotoxins that are produced in pure culture by the fungal species listed in Tables 3 and 4 are given. Of all of the species in Tables 3 and 4, only Alt. infectoria species-group and P. solitum are regarded as non-toxigenic. As can been seen from Table 5, several fungal species within the same genus and found in the same product can produce the same mycotoxins (e.g. roquefortine C and penitrem A by Penicillia in fruit or culmorins and trichothecenes by Fusaria in cereals). In fruit that has either been damaged in the orchard and/or stored poorly, one or more of the following toxic metabolites might theoretically be found: altenuene, alternariols, chaetoglobosins, citrinin, patulin, roquefortine C and tenuazonic acid. In fresh cereals that have been harvested during a rainy period, the following toxic metabolites would be relevant: antibiotic Y, beauvericin, culmorins, enniatins, fusarin C, fusarochromanone, moniliformin, trichothecenes and zearalenone. In cereals that have been stored poorly and/or not dried down after harvest, the following toxic metabolites should be considered: aflatoxins, aspergillic acid, citrinin, cyclopiazonic acid, nephrotoxic glycopeptides, ochratoxin A, penicillic acid, terphenyllin, verrucosidin, viomellein, vioxanthin, viridic acid, xanthoascin and xanthomegnin. However, it should be noted that mycotoxins produced in the field or at an early stage of storage always should be taken into consideration, as mycotoxins in general are persistent through storage and processes for food and feed production.

3.5.

Production of Toxic Metabolites in Fruit and Cereals

Analyses of extracts from pure fungal cultures can indicate which fungal metabolites should be analysed, but the mycobiota of the actual sample needs to be determined to make realistic recommendations. Mycotoxins that have been detected in naturally moulded apples, cherries, barley and wheat samples examined in our laboratory are given in Table 6. Samples that were mouldy when received were extracted immediately, while sound samples were incubated for a week

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Table 5. Fungal species found in apples, cherries wheat and barley from the north temperate zone and some mycotoxins they are known to produce Toxigenic fungal species Mycotoxins produced in pure culture Alternaria arborescens sp.-grp. Altenuene, alternariols, altertoxins, tenuazonic acid Alt. tenuissima sp.-grp. Altenuene, alternariols, altertoxins, tenuazonic acid Aspergillus candidus Terphenyllin, xanthoascin Asp. flavus Aflatoxin, aspergillic acid, cyclopiazonic acid Asp. niger Malformins, naphtho-γ-pyrones Bipolaris sorokiniana Sterigmatocystin Byssochlamys spp. Byssochlamic acid, patulin Fusarium avenaceum Antibiotic Y, aurofusarin, enniatins, fusarin C, moniliformin F. culmorum Aurofusarin, culmorin, fusarin C, trichothecenes, zearalenone F. equiseti Fusarochromanone, trichothecenes, zearalenone F. graminearum Aurofusarin, culmorin, fusarin C, trichothecenes, zearalenone F. langsethiae Culmorin, enniatins, trichothecenes F. lateritium Antibiotic Y, enniatins, F. poae Beauvericins, culmorins, fusarin C, trichothecenes F. sporotrichioides Aurofusarin, beauvericins, culmorins, fusarin C, trichothecenes F. tricinctum Antibiotic Y, aurofusarin, enniatins, fusarin C, moniliformin Penicillium aurantiogriseum Penicillic acid, verrucosidin, nephrotoxic glycopeptides P. carneum Patulin, isofumigaclavin, penitrem A, roquefortine C P. crustosum Penitrem A, roquefortine C P. cyclopium Penicillic acid, xanthomegnin, viomellein, vioxanthin P. expansum Citrinin, chaetoglobosins, communesins, patulin, roquefortine C P. freii Penicillic acid, xanthomegnin, viomellein, vioxanthin P. hordei Roquefortine C P. melanoconidium Penicillic acid, penitrem A, xanthomegnin P. polonicum Penicillic acid, verrucosidin, nephrotoxic glycopeptides P. verrucosum Citrinin, ochratoxin A P. viridicatum Penicillic acid, viridic acid, xanthomegnin, viomellein

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Table 6. Mycotoxins detected in naturally infected samples Sample Metabolites detected Immature apples with mouldy core (1) Alternariols, antibiotic Y, aurofusarin Immature apples with mouldy core (2) Altenuene, alternariols Mature apples with mouldy core (3) Alternariols, patulin Mouldy apple pulp (4) Alternariols, antibiotic Y, aurofusarin, ascladiol, Mouldy apple on tree (5) Chaetoglobosin A Mouldy sweet cherries ‘June drop’ (6) Alternariols, antibiotic Y Mouldy, mature sweet cherries (7) Alternariols Mouldy cherry juice (8) Chaetoglobosins, communesins, roquefortine C Mouldy barley (9) Ochratoxin A Mouldy barley ‘hot spot’ (10) Ochratoxin A Mouldy wheat (11) Antibiotic Y, ochratoxin A, zearalenone Mouldy wheat (12) Aurofusarin, fusarin C, zearalenone

or until mould could be seen to simulate a worst case. Fusarium and/or Alternaria metabolites were detected in samples of immature apples from the field with visible fungal growth (samples 1 and 2, worst cases), while the mature, fresh apple samples (samples 3 and 4, worst cases) also contained Penicillium metabolites. One mature, mouldy apple (sample 5) still hanging on the tree contained only chaetoglobosin A. The cherry sample (sample 6), which consisted of immature cherries that had been shed by the trees, had 100 % infection with Alternaria species. Extraction showed that it contained high amounts of both Alternaria and Fusarium metabolites, whereas the mature cherries (sample 7, worst case) only contained Alternaria metabolites. The mouldy cherry juice (sample 8), on the other hand, contained only Penicillium metabolites. The mouldy barley samples (samples 9 and 10), which had been stored without drying after harvest, contained ochratoxin A. Sample 10, sampled in the ‘green hot spot’ of the mouldy lot, contained approximately 1000 times the amount of ochratoxin A as sample 9, which had little visible fungal growth. In the mouldy wheat sample (sample 11) Fusarium metabolites as well as ochratoxin A were detected, while sample 12 only contained Fusarium metabolites.

4.

DISCUSSION

Analyses of seasonal changes in apple mycobiota showed that many fungal species found in mature fruit already were present in the flowers and later colonized the immature fruit (Table 1). Spraying with

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fungicides decreased Botrytis infection and had some effect on Cladosporium numbers, but no effect on Alternaria, Fusarium and Penicillium. The number of Alternaria, Fusarium and Penicillium infected apples also increased after spraying and continued to increase until apples were picked. In cherries, the same fungal species were seen in mature cherries as in flowers (Table 1). Application of fungicides had an effect on all the fungal genera found in flowers; Alternaria infections decreased by 50-65%. Alternaria and Cladosporium numbers, however, increased again before harvest in sour cherries, while the numbers of Alternaria remained constant in sweet cherries, probably due to the early drop of immature cherries, which had 100% infection with Alternaria. Our results show that the mycobiota of apples and cherries are similar at genus level, but different in species composition. Alternaria tenuissima species-group, P. expansum and P. solitum dominate in apples, whereas A. arborescens species-group, and Stemphylium spp. dominate in cherries (Table 3). Analyses of the mycobiota in cereals from sowing to harvest showed, in contrast to fruit, that the initial mycobiota present in the untreated seed played only a small role in the subsequent mycobiota on mature kernels, though it may play a great role in the viability of the seed (Table 2). The two untreated seed samples contained a high number of Alternaria, Penicillium and Eurotium species. Surface disinfection removed 80-90% of the Penicillium and Eurotium numbers, while the same only could be done for 40-45% of the Alternaria. Furthermore, the Penicillium species in the seed and in the immature kernels were different. In the seed P. chrysogenum, P. cyclopium and P. freii were found, whereas P. aurantiogriseum, P. polonicum and P. verrucosum were found in mature kernels growing from the untreated seed. The only fungi that were found in larger amounts in seed and recovered in more than 50% of the harvested cereal samples belonged to A. infectoria species-group. The numbers of Alternaria and Fusarium found in the three mature samples were low (less than 60%) and high (more than 80%), respectively, compared with other reports (Andersen et al., 1996; Kosiak et al., 2004). A very wet period in June and July 2002, during the growing season in Western Denmark was probably responsible. Comparisons of the mycobiota from two mature barley samples grown from fungicide treated and untreated seed showed few differences. As fruits mature and are harvested, fungi such as Botrytis, Monilia and Zygomycetes are known to cause fruit spoilage in orchards as well as in storage, whereas Cladosporium and Epicoccum are known for their discolouration of cereals in the field. These fungi cause economical losses, but none of them are associated with production of mycotoxins.

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Different species of Alternaria, Fusarium and Penicillium, on the other hand, all spoil fruit and cereals, but produce species specific mycotoxins (Table 5) and hundred mycotoxins and other biologically active metabolites from these three genera have been characterised within recent years (Nielsen and Smedsgaard, 2003) and it is reasonable to expect that more than the few included in the legislation (aflatoxin, ochratoxin, deoxynivalenol, fumonisins in cereals and patulin in fruits) can be produced in mouldy foods. The results presented in this study show that Alternaria and Fusarium in fruit and cereals may pose a mycotoxin risk. During spoilage of apples and cherries, P. expansum is known to produce patulin, which has been incorporated in the legislation on fruit produce. However, both Alternaria and Fusarium were able to produce additional metabolites in mouldy fruit samples (Table 6, sample 4): alternariols, antibiotic Y and aurofusarin. In cereals, P. verrucosum is known to produce ochratoxin A, which has also been incorporated in the legislation on raw cereal grain. However, Fusarium was able to produce antibiotic Y and zearalenone in addition to ochratoxin A from P. verrucosum in mouldy wheat (Table 6, sample 11). For these lesser known metabolites no or very limited data are available on the toxicity on co-produced metabolites and their possible synergistic effects, which make risk assessment in food and food production systems difficult. In conclusion, we see the co-occurrence of these specific Alternaria and Fusarium metabolites and their potential toxicities as the major future challenge in food mycology.

5.

ACKNOWLEDGEMENTS

The authors are grateful to Dr. Jens C. Frisvad for discussion of manuscript and identification of some of the Penicillium cultures. This work was partly supported by the Danish Ministry of Food, Agriculture and Fisheries through the program “Food Quality with a focus on Food Safety”, by LMC Centre for Advanced Food Studies and by the Danish Technical Research Council through ‘Program for Predictive Biotechnology’.

6.

REFERENCES

Abildgren, M. P., Lund, F., Thrane, U., and Elmholt, S., 1987, Czapek-Dox agar containing iprodione and dicloran as a selective medium for the isolation of Fusarium species, Lett. Appl. Microbiol. 5:83-86.

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Andersen, B., Thrane, U., Svendsen, A., Rasmussen, I. A., 199, Associated field mycobiota on malting barley, Can. J. Bot. 74:854-858. Andersen, B., Krøger, E., and Roberts, R.G., 2002, Chemical and morphological segregation of Alternaria arborescens, A. infectoria and A. tenuissima speciesgroups, Mycol. Res. 106:170-182. Andersen, B., Smedsgaard, J., and Frisvad, J. C., 2004, Penicillium expansum: consistent production of patulin, chaetoglobosins and other secondary metabolites in culture and their natural occurrence in fruit products, J. Agric. Food Chem. 52:2421-2428. Burgess, L. W., Summerell, B. A., Bullock, S., Gott, K. P., and Backhouse, D., 1994, Laboratory Manual for Fusarium Research. 3rd Edition, University of Sydney, Sydney, Australia. EC (European Commission), 2004, European Commission Regulation 455/2004 of 11 March, 2004. European Commission, http://europa.eu.int/eur-lex/pri/en/oj/dat/2004/ l_074/l_07420040312en00110011.pdf Filtenborg, O., Frisvad, J. C., and Thrane, U., 1996, Moulds in food spoilage, Int. J. Food Microbiol. 33:85-102. Frisvad, J. C., 1983, A selective and indicative medium for groups of Penicillium viridicatum producing different mycotoxins on cereals, J. Appl. Bacteriol. 54:409-416. Frisvad, J. C., Thrane, U., and Filtenborg, O., 1998, Role and use of secondary metabolites in fungal taxonomy, in: Chemical Fungal Taxonomy, J. C. Frisvad, P. D. Bridge, and D. K. Arora, (eds), Marcel Dekker, New York, pp. 289-319. Gerlach, W., and Nirenberg, H., 1982, The genus Fusarium -A Pictorial Atlas, Mitteilungen aus der Biologische Bundesanstalt für Land-und Forstwirtschaft, Berlin-Dahlem 209:1-406. Hocking, A. D., and Pitt, J. I., 1980, Dichloran-glycerol medium for enumeration of xerophilic fungi from low moisture foods, Appl. Environ. Microbiol. 39:488-492. Kosiak, B., Torp, M., Skjerve, E., and Andersen, B., 2004, Alternaria and Fusarium in Norwegian grains of reduced quality -a matched pair sample study, Int. J. Food Microbiol. 93:51-62. Marasas, W. F. O., Nelson, P. E., and Toussoun, T. A., 1984, Toxigenic Fusarium Species. Identity and Mycotoxicology, The Pennsylvania State University Press, University Park, pp. 1-328. Nielsen, K. F., and Smedsgaard, J., 2003, Fungal metabolite screening: database of 474 mycotoxins and fungal metabolites for de-replication by standardised liquid chromatography-UV detection-mass spectrometry methodology, J. Chromatogr. A. 1002:111-136. Nirenberg, H., 1976, Untersuchungen über die morphologische und biologische Differenzierung in der Fusarium-Sektion Liseola, Mitteilungen aus der Biologische Bundesanstalt für Land-und Forstwirtschaft. Berlin-Dahlem 169:1-117. Nirenberg, H. I., 1989, Identification of Fusaria occurring in Europe on cereals and potatoes, in: Fusarium: Mycotoxins, Taxonomy and Pathogenicity, J. Chelkowski (ed.), Elsevier Science Publishers B.V., Amsterdam, pp. 179-193. Pitt, J. I., and Hocking, A. D., 1997, Fungi and Food Spoilage, Blackie Academic and Professional, London, pp. 1-593. Samson, R. A., Frisvad, J. C., 2004, Penicillium subgenus Penicillium: new taxonomic schemes, mycotoxins and other extrolites, Stud. Mycol. 49:1-257. Samson, R. A., Hoekstra, E. S., Frisvad, J. C., (eds), 2004, Introduction to Foodand Airborne Fungi. 7th Edition. Centraalbureau voor Schimmelcultures, Utrecht, pp. 1-389.

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Simmons, E. G., 1967, Typification of Alternaria, Stemphylium and Ulocladium, Mycologia 59:67-92. Simmons, E. G., 1969, Perfect states of Stemphylium, Mycologia 61:1-26. Simmons, E. G., 1986, Alternaria themes and variations (22-26), Mycotaxon 25:287-308. Simmons, E. G., 1992, Alternaria taxonomy: Current Status, viewpoint, challenge, in: Alternaria Biology, Plant Diseases and Metabolites, J. Chelkowski and A. Visconti, eds, Elsevier, Amsterdam, pp. 1-35. Simmons, E. G., and Roberts, R. G., 1993, Alternaria themes and variations (73), Mycotaxon 48:109-140. Smedsgaard, J., 1997, Micro-scale extraction procedure for standardized screening of fungal metabolites production in cultures, J. Chromatogr. A 760:264-270. Thrane, U., 2001, Developments in the taxonomy of Fusarium species based on secondary metabolites, in: Fusarium. Paul E. Nelson Memorial Symposium. B. A. Summerell, J. F. Leslie, D. Backhouse, W. L. Bryden, and L. W. Burgess (eds), APS Press, St. Paul, Minnesota, pp. 29-49.

BLACK ASPERGILLUS SPECIES IN AUSTRALIAN VINEYARDS: FROM SOIL TO OCHRATOXIN A IN WINE Su-lin L. Leong,*‡ Ailsa D. Hocking,* John I. Pitt,* Benozir A. Kazi,† Robert W. Emmett† and Eileen S. Scott‡§

1.

INTRODUCTION

Fungi classified in Aspergillus Section Nigri (the black Aspergilli) are ubiquitous saprophytes in soils around the world, particularly in tropical and subtropical regions (Klich and Pitt, 1988; Pitt and Hocking, 1997). Several species in this Section are common in vineyards and are often associated with bunch rots (Amerine et al., 1980). A. niger is reported to be the primary cause of Aspergillus rot in grapes before harvest (Nair, 1985; Snowdon, 1990), while A. aculeatus (Jarvis and Traquair, 1984) and A. carbonarius (Gupta, 1956) have also been reported. The development of fungal bunch rots has been correlated with the splitting of grape berries (Barbetti, 1980), and Aspergillus counts on grapes grown for drying were greater during seasons when rain before harvest caused the berries to split (Figure 1) (Leong et al., 2004). Spores of black Aspergillus spp. are resistant to UV light (Rotem and Aust, 1991), which may account for their

* Su-lin L. Leong, Ailsa D. Hocking and John I. Pitt, CSIRO Food Science Australia, North Ryde, NSW 2113, Australia. † Benozir A. Kazi and Robert W. Emmett, Department of Primary Industries, Mildura, Victoria 3502, Australia. ‡ Su-lin L. Leong and Eileen S. Scott, University of Adelaide, Glen Osmond, South Australia 5064, Australia. § All authors, Cooperative Research Centre for Viticulture, Glen Osmond, South Australia 5064, Australia. Correspondence to: [email protected]

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A. aculeatus A. carbonarius

800,000

A. niger

600,000 400,000 200,000 0 1998

1999

2000

Year

Figure 1. Mean severity of infection of fresh and drying grapes (combined) by each of three black Aspergillus species over three successive harvest seasons. Rain occurred before harvest in 1999 and 2000. The mean count was derived by summing the counts for each species of black Aspergillus from all the fruit samples, and dividing that sum by the total number of samples. Reproduced from Leong, S. L., Hocking. A. D., and Pitt, J. I., 2004, Australian Journal of Grape and Wine Research 10: 83-88 (with permission from the Australian Society of Viticulture and Oenology).

persistence in vineyards and on grape berries even after drying (King et al., 1981; Abdel-Sater and Saber, 1999; Abarca et al., 2003). Within Section Nigri, A. carbonarius and A. niger have been shown to produce the mycotoxin, ochratoxin A (OA) (Abarca et al., 1994; Téren et al., 1996; Heenan et al., 1998; reviewed in Abarca et al., 2001). OA is a demonstrated nephrotoxin, which may also be carcinogenic, teratogenic, immunogenic and genotoxic. It has been classified as Group 2B, a “possible human carcinogen” (Castegnaro and Wild, 1995). OA has been detected in grapes and grape products including juice, wine, dried vine fruit and wine vinegars (Zimmerli and Dick, 1996; MacDonald et al., 1999; Majerus et al., 2000; Markarki et al., 2001; Da Rocha Rosa et al., 2002; Sage et al., 2002; OA in wine and grape juice reviewed by Bellí et al., 2002). A survey of 600 Australian wines showed that OA was present only at low levels. Only 15% of samples had levels > 0.05 µg/l and 85% of these were < 0.2 µg/l. The maximum level found was 0.61 µg/l (Hocking et al., 2003). In Europe, a limit of 2 µg/kg in table wines is under discussion (Anon., 2003) and a limit of 10 µg/kg in dried vine fruits has been set (European Commission, 2002). OA in grapes and grape products is produced by toxigenic A. carbonarius and A. niger species which have been isolated from grapes in

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France (Sage et al., 2002), South America (Da Rocha Rosa et al., 2002), Spain (Cabañes et al., 2002), Italy (Battilani et al., 2003b), Portugal (Serra et al., 2003) and Greece (Tjamos et al., 2004). In an extensive study of Australian dried vine fruit, strains of A. carbonarius were commonly isolated from semi-dried and dried vine fruit in the field, and all were capable of producing OA in the laboratory (Leong et al., 2004). Hence A. carbonarius is thought to be the primary species responsible for OA production in grapes in Australia. Assuming that OA production in grapes ceases at the commencement of processing, typically a sterilisation step in industrial juice and wine production (Roset, 2003), the concentration of OA in the final product is a function of the initial concentration in the grapes and the effect of processing. Processes which reduce OA can be classified into two groups, physical removal and degradation. Physical removal of OA first involves removing the site where OA has been produced, for example, the removal of visibly mouldy berries from table grapes. It is not well understood if OA is primarily associated with the skin, pulp or juice of grape berries. However, a strong association with the skin or pulp would suggest that a relatively small proportion of OA remains in the finished beverage. The high water content of grape berries may lead to the migration of OA from the zone of fungal growth to other parts of the berry (Engelhardt et al., 1999). A second aspect of physical removal of OA is the partitioning of the toxin between solid and liquid phases during processing. Fernandes et al. (2003) conducted microvinification trials on crushed grapes spiked with OA, and reported reductions in OA of 50-95%. The most significant reductions resulted from solid-liquid separation steps, such as pressing the juice or wine from the skins, and decanting the wine from precipitated solids. Many of the solids present in grape juice have an affinity for OA and will loosely bind and precipitate the toxin from solution (Roset, 2003), as do some fining agents added during winemaking, such as activated charcoal (Dumeau and Trione, 2000; Castellari et al., 2001; Silva et al., 2003). Little is known about the degradation of OA by wine yeasts during fermentation, though this has been demonstrated during beer fermentation (Baxter et al., 2001). Silva et al. (2003) reported reduction in OA by lactic acid bacteria during malolactic fermentation which follows the completion of primary (yeast) fermentation. However, Fernandes et al. (2003) argued that this is not a true degradation, rather, bacterial biomass binding OA that later settles out of the wine. The addition of sulphur dioxide and the pasteurisation of juice by heating have no effect on OA (Roset, 2003).

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This paper presents original data on OA contamination of wine, covering the source of A. carbonarius in Australian vineyards, the survival of A. carbonarius spores on the surface of bunches, and the passage of OA throughout vinification of grapes inoculated with A. carbonarius.

2.

MATERIALS AND METHODS

2.1.

Aspergillus carbonarius in the Vineyard Environment

Substrates were collected from vineyards in the grape-growing region centred around Mildura, Victoria, Australia. Substrates collected included parts of vines [green, yellow (senescing) and dead leaf tissue, green and dead berries, dead bunch remnants (dried rachides), tendrils, canes and bark] and materials from the vineyard floor [green cover crop plants, dead cover crop trash, vine trash and soil]. Collections were made over three growing seasons, from six vineyards in 2000-01 and from three vineyards in 2001-02 and 2002-03. In 200001, samples of substrates were collected from three sites along a diagonal transect in each vineyard 2 weeks after veraison and at harvest. In the latter two seasons, samples of substrates were collected from five sites along a diagonal transect in each vineyard when berries were pea size, at 2 weeks after veraison and at harvest. To quantify A. carbonarius present on the surface of these substrates, samples were washed for 2 min in sterile water containing Citowett® (BASF Australia Ltd, Victoria, Australia) as a wetting agent, and aliquots of the solution were plated in duplicate onto Dichloran Rose Bengal Chloramphenicol Agar (DRBC) (Pitt and Hocking, 1997). Serial dilutions were performed on soil samples, followed by plating onto DRBC. After plates were incubated at 25°C for 5-7 days, colonies of A. carbonarius were identified and enumerated. The presence of A. carbonarius in vineyard soils was also compared among four vineyards in 2002-03 by dilution plating as described above. Thirty soil samples were collected from each vineyard, ten samples at each stage of vine growth, i.e. when berries were pea size, at 2 weeks after veraison and at harvest. The tillage practices of the vineyards were noted. Soil was sampled directly under vines and also between vine rows. In one vineyard, five soil sampling cores of 0.2 m2 were taken from under vines. For each soil core, A. carbonarius was enumerated in soil at the surface, 5 cm and 15 cm below the surface. A. carbonarius was also enumerated in the air of this vineyard. Colonies from 20 l of air sampled using a MAS-100® air sampler

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(Merck KGaA, Darmstadt, Germany) were enumerated on DRBC. Samples were taken on nine occasions from air in the vineyard at 10 cm, 100 cm and 180 cm above the vineyard floor. Statistical analyses were performed using Genstat (6th edition, Lawes Agricultural Trust, Rothamsted, UK).

2.2.

Survival of Aspergillus carbonarius Spores on the Surface of Bunches Preharvest

A trial was conducted in the Hunter Valley, New South Wales, Australia to examine the survival of A. carbonarius spores on the surface of Chardonnay and Shiraz grapes (three replicate rows) during the growing season in 2002-03. The vines were over 25 years old, trained onto horizontal wires and under drip irrigation. Spores of 7-14 day old cultures of A. carbonarius on Czapek Yeast Agar (CYA) (Pitt and Hocking, 1997) plates were harvested into sterile water containing Tween-80® (0.05% w/v; Merck, Victoria, Australia), and diluted to 2-4 × 105 colony forming units per ml (cfu/ml). Bunches were inoculated by immersion in 1 l of suspension contained within a plastic bag. The same inoculum was used for up to 40 bunches without decrease in the spore concentration. Twelve bunches in each row were inoculated at pre-bunch closure (berries green and pea size), veraison and 11-16 days preharvest. Two bunches were combined into a single sample, resulting in six samples per replicate at each sampling stage. Inoculated bunches were sampled after the inoculum had dried to give an initial value, at each of the subsequent stages and at harvest. Bunches were homogenised for 3 min in a stomacher (BagMixer, Interscience, France) with the addition of sterile distilled water, and serial dilutions of the suspension were plated onto DRBC. After incubation for 3 days at 25°C, colonies of A. carbonarius were enumerated. The average berry weight at each growth stage was calculated, and the number of A. carbonarius colonies was expressed as cfu per berry, in order to compare the number of viable spores present during each stage.

2.3.

Winemaking

2.3.1.

Inoculation and Vinification of Grapes

Berries were inoculated preharvest with a spore suspension of A. carbonarius (prepared as described in 2.2) at approximately 107 cfu/ml.

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Strains selected for inoculation were local to the region of the experimental vineyard, and were strong producers of OA when screened on Coconut Cream Agar (CCA) (Heenan et al., 1998). A variety of inoculation techniques was employed, all involving puncture damage to the berry skin and subsequent contact with the spore suspension. In addition to the primary inoculation, a supplementary inoculation of additional fruit was often performed towards harvest to ensure sufficient fruit for vinification. At harvest, inoculated and uninoculated fruit were mixed to simulate high, intermediate and low or absent levels of OA in fruit. Table 1 summarises the inoculation, incubation and harvest details.

Table 1. Preparation of OA-contaminated grapes for winemaking Location, Mildura, Victoria, 2002 Hunter Valley, New South vintage Wales, 2003 A. carbonarius FRR 5374a, FRR 5573, FRR 5682, FRR 5683 strains FRR 5574 Grape variety Chardonnay Shiraz Semillon Shiraz Method of Berries injected Berries Berries Berries inoculation using syringe injected using punctured injected {berries syringe {skin with a bed of using injected scored using pins dipped syringe using syringe}b grater and in spore sprayed with suspension spore {berries suspension} injected using syringe} Period from 21 days 14 days 9 days 8 days primary inoc. {4 days}b {13 days} {3 days} until harvest High OA wine: 53 kg 120 kg 25 kg 28 kg mass of grapes inoculated inoculated inoculated inoculated Intermediate 34 kg 46 kg 15 kg 11 kg OA wine: mass inoculated inoculated inoculated inoculated of grapes + 23 kg + 73 kg + 13 kg + 16 kg uninoculated uninoculated uninoculated uninoculated Control wine: 51 kg 118 kg 32 kg 27 kg mass of grapes uninoculated uninoculated uninoculated uninoculated Size of 4l 16 l 2l 4l ferment including including skins skins a

FRR numbers are from the culture collection of Food Science Australia, North Ryde, NSW, Australia. b {} bracketed text refers to supplementary inoculation of additional fruit.

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Harvested bunches were chilled at 4°C prior to crushing. After crushing, eight samples of must from each toxin level were collected in order to establish the initial total OA present in the berries. Samples were also collected throughout the vinification process as described below. During white vinification, the must was pressed, after which potassium metabisulphite was added to give 50 ppm SO2 in the juice. Pectinase was added in the form of Pomolase AC50 (0.05 ml/l juice; Enzyme Solutions, Victoria, Australia) or Pectinase (0.5 g/l juice; Fermtech, Queensland, Australia). The juice was overlaid with nitrogen or carbon dioxide, and refrigerated at 4°C for at least 24 h to precipitate solids. In 2002, the juice was divided into four replicate ferments at each toxin level before clarification; this division occurred after clarification in 2003. The pH was adjusted to approximately 3.3 by the addition of tartaric acid to give a titratable acidity of 6.5-7.0 g/l. The clarified juice was siphoned into bottles filled with nitrogen or carbon dioxide and fitted with stoppers and air traps. The yeast QA23 (Lallemand, Toulouse, France) was rehydrated and added at a rate equivalent to 0.2 g dry yeast/l juice. Diammonium phosphate was added at 0.5 g/l juice. The fermentation temperature was 19°C in 2002 and 15°C in 2003. Diammonium phosphate was added during fermentation as required and after fermentation was completed, the wine was racked. Potassium metabisulphite was added at a rate equivalent to 50 ppm SO2 to stabilise the wine and prevent further fermentation. Bentonite (0.5 g/l; Fermtech, Queensland, Australia) and Liquifine (2002: 1 ml/l; 2003: 0.6 ml/l; Winery Supplies, Victoria, Australia) were added, and the bottles placed at 19°C (2002) or 15°C (2003) to allow precipitation of solids. A second racking was performed for all bottles, and potassium metabisulphite was added to bring the free SO2 to 20 ppm. The bottles were placed at < 4°C for cold stabilisation for > 30 days. The wine was filtered into 375 ml glass bottles with cork closures. During red vinification, the must was divided into 4 replicate fermentations at each toxin level. Potassium metabisulphite was added to give 50 ppm SO2 in the must, diammonium phosphate was added at 0.5 g/l must, and tartaric acid was added to bring the titratable acidity to 6.5 g/l. The yeast D254 (Lallemand, Toulouse, France) was rehydrated and added at approximately 0.3 g/l must. The cap was plunged 2-3 times daily. The must was pressed after 4 days of fermentation at room temperature in 2002, and after 6 days of fermentation at approximately 20°C in 2003. Fermentation was finished in bottles at room temperature. During the first racking, 50 ppm SO2 was added, after

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which the wine was held at 19°C (2002) or 15°C (2003) to precipitate yeast cells and other solids. At the second racking, SO2 was added to maintain a final concentration of 50 ppm. The wine was held at < 2°C for cold stabilisation, after which the pH was adjusted and the wine bottled through a filtration line. Bottles were cellared at room temperature (approximately 22°C) in 2002, and at 15°C in 2003. 2.3.2.

OA Assays

A new method was developed for the rapid analysis of OA in grape matrices. Samples were standardised by weight. Grape musts were homogenised and a 10 g subsample weighed into a centrifuge tube. Methanol (10 ml), Milli-Q water (10 ml) and 10N HCl (≈ 0.15 ml) were added, and mixed thoroughly with the sample. For liquid samples, 10 g of sample was mixed with methanol (1.5 ml) and 10N HCl (≈ 0.15 ml). The mixture was centrifuged at 2500 rpm for 15 min. A 900 mg C18 solid phase extraction cartridge (MaxiClean, Alltech, Deerfield, USA) was conditioned with 5 ml acetonitrile followed by 5 ml water, and the supernatant was passed dropwise through this cartridge under vacuum. The pellet was resuspended in 10 ml 10% methanol, then centrifuged for a further 15 min at 2500 rpm. This supernatant was also passed through the C18 cartridge. For must samples, an additional 10 ml water was washed through the C18 cartridge at this stage. A 200 mg aminopropyl cartridge (4 ml Extract-Clean, Alltech, Deerfield, USA) was conditioned with 3 ml methanol. The C18 and aminopropyl cartridges were attached in series, and the sample was eluted from the C18 cartridge onto the aminopropyl cartridge with the addition of 10 ml methanol. The sample was eluted from the aminopropyl cartridge with 10 ml 35% ethyl acetate in cyclohexane containing 0.75% formic acid. The eluate was dried under reduced pressure at 45°C and was resuspended in 1 ml mobile phase (35% acetonitrile containing 0.1% acetic acid) for analysis by HPLC (Hocking et al., 2003). Aliquots of the wine extracts were chromatographed on an Ultracarb (30) C18 4.6 × 250 mm, 5 µm column (Phenomenex, Torrance, USA). The mobile phase consisted of acetonitrile: water:acetic acid (50:49:1, v/v), and was delivered through the heated column (40°C) at a flow of 1.3 ml/min using a Shimadzu 10A VP high pressure binary gradient solvent delivery system. Detection of OA was achieved by post column addition of ammonia (12.5% w/w,

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0.2 ml/min) and monitoring the natural fluorescence of OA at 435 nm after excitation at 385 nm (Shimadzu, RF-10AXL). Sample injections were performed using a Shimadzu SIL-10Advp autosampler, and the typical injection volume was 30 µl. OA in the wine extracts was quantified by comparison with a calibration curve. Typical recoveries ranged from 80-100%. The results presented have not been corrected for recovery.

3.

RESULTS AND DISCUSSION

3.1.

Aspergillus carbonarius in the Vineyard Environment

Counts of A. carbonarius were high in soil and in vine trash on soil and relatively low on other substrates (Table 2). Hence, soil is likely to be the primary source of A. carbonarius in vineyards. In the four vineyards surveyed, soil beneath vines contained more A. carbonarius than soil between rows (P < 0.05, Figure 2). This association is likely to be due to damaged and dead berries falling onto the soil and providing a sugar-rich medium for the growth of indigenous saprophytic Aspergillus species. The concentration of A. carbonarius propagules was highest at the surface of soil, where this debris is found, and decreased deeper within the soil profile of an untilled vineyard (Figure 3). The vineyard in which the soil profile was regularly disturbed by tilling contained a higher concentration of A. carbonarius in the soil than vineyards in which the soil was less disturbed (P < 0.05, Figure 2). In vineyards with minimal tillage, A. carbonarius may be one member of a complex and stable microbial community associated with the cover crop and other flora on the soil surface. One potential effect of regular tillage is to allow the increase of a dominant species (Marfenina and Mirchink, 1989). Table 2. Aspergillus carbonarius on vineyard materials. Results expressed as cfu/ml surface wash unless otherwise indicated. Substrate 2000-01 2001-02 2002-03 Canes (dead) 8 1 not recorded Vine bark 31 9 not recorded Bunch remnants 47 15 not recorded Cover trash (dead) 8 10 2 Vine trash on soil 669 45 20 Soil (cfu/g) 4,987 1,219 1,342

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4,000

Between rows Under vines

B

3,000

2,000

A

A

b

1,000 a

a

a

0 1

2

3

4

Vineyard

Figure 2. Aspergillus carbonarius in soil under vines and between vine rows in vineyards with minimal and continual tillage, n = 30. Vineyard 1 and 3: No tillage for the last 3 and 4 years, respectively. Vineyard floors were covered with wild grasses and/or weeds. Vineyard 2: Tilled once each year before sowing a rye grass cover crop. Vineyard 4: Tilled monthly after a cover crop was established. Vertical bars with different letters differ significantly (LSD, P < 0.05).

A. carbonarius may compete more effectively for sugars from fallen berries, and tillage would also distribute propagules throughout the soil profile. Other authors have observed higher levels of some fungi in tilled soil than in untilled soil. An example is the primary 1,500 Mean A. carbonarius count (cfu/g soil)

b 1,000

a

500

a 0

0-1 cm

5 cm Depth

15 cm

Figure 3. Aspergillus carbonarius in untilled vineyard soil at different depths in 2001-02, n = 5. Vertical bars with different letters differ significantly (LSD, P < 0.05).

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pathogen of root rot of wheat, Cochliobolus sativus (Diehl, 1979; Duczek, 1981; Diehl et al., 1982). A. carbonarius was present in vineyard air, though the concentration of conidia decreased with height from the vineyard floor (Figure 4). This suggests that wind may distribute conidia of A. carbonarius from the soil onto berry surfaces. During the early stages of berry development from pre-bunch closure until veraison, A. carbonarius spores survived poorly and a nine fold decrease in the number of viable propagules was observed (Figure 5). This suggests that the surface of green berries is a hostile environment for the survival of spores. The vine canopy during the early part of the season in 2002-2003 was sparse due to drought. Thus, the berries were not shielded from UV light by overhanging leaves. A sparse canopy would also allow greater penetration of routine fungicide sprays, which, together with the UV light may have contributed to the death of the spores (Rotem and Aust, 1991).

3.2.

Survival of Aspergillus carbonarius Spores on Bunch Surfaces Preharvest

For Shiraz bunches inoculated at pre-bunch closure, there was a consistent decrease in the counts of A. carbonarius between the inoculation time and veraison. This decrease continued in some bunches between veraison and harvest (Figure 6). However, in other bunches, the count increased due to infection of berries and subsequent

Mean A. carbonarius count (cfu/L air)

0.8

c

0.6

0.4

b a

0.2

0 10 cm

100 cm

180 cm

Height

Figure 4. Aspergillus carbonarius in air at different levels above the vineyard floor, n = 9. Vertical bars with different letters differ significantly (LSD, P < 0.05).

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3.25

A. carbonarius count (log cfu per berry)

3 2.75 2.5 2.25 2 1.75 1.5 Pre-bunch closure

Veraison

A. carbonarius count (cfu per berry)

Figure 5. Spore death on berry surfaces between pre-bunch closure and veraison; n = 32, comprising both Chardonnay and Shiraz varieties. 50% of the data are contained within the boxes, while the bar within the box plot shows the median value. Maximum and minimum values are indicated by vertical lines. The difference between Aspergillus carbonarius spore count per berry at pre-bunch closure and veraison was significant at P < 0.001.

Pre-bunch closure, day 0 Veraison, day 27 Preharvest, day 62 Harvest, day 78

1,000,000 100,000 10,000 1,000 100 10 1 Row A

Row B

Row C

Figure 6. Survival of Aspergillus carbonarius inoculated on the surface of Shiraz berries at pre-bunch closure.

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sporulation by the mould. A slight but statistically significant decrease in the counts of A. carbonarius was also observed on Chardonnay bunches inoculated 11 days before harvest (Figure 7). There were no fungicide sprays applied during this period, hence death of spores could be attributed to residual fungicide activity and/or exposure to UV light. Samples with increased A. carbonarius counts had foci of infection and sporulation on some berries. These results suggest that the critical time for the development of Aspergillus rots occurs from veraison onwards. Battilani et al. (2003a) and Serra et al. (2003) both noted black Aspergillus spp. were more frequently isolated from berries after veraison.

3.3.

Winemaking

HPLC analysis of samples taken throughout vinification showed that the greatest reduction in OA was observed at pressing, as the mean concentration of OA in white juice was 26% of the concentration in crushed grapes (must); the corresponding figure for Shiraz was 28% (Figure 8). This suggests that there is a strong association of OA with the skins and seeds (marc) trapped during pressing. Clarification of white juice with pectinase and precipitation of solids overnight resulted in a mean reduction in OA of an additional 12%, within the range observed for precipitation of must sediments during industrial

A. carbonarius count (cfu per berry)

100,000

Preharvest, day 0 Harvest, day 11

10,000

1,000

100

10

1 Row A

Row B

Row C

Figure 7. Survival of Aspergillus carbonarius inoculated on the surface of Chardonnay berries preharvest. The difference between A. carbonarius spore count per berry at preharvest and harvest was significant at P < 0.05.

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Proportion of OA relative to initial concentration in grapes

100% 80% 60% 40% 20% 0% Chard, intermed (2002)

Chard, high (2002)

Sem, intermed (2003)

Sem, high (2003)

Variety, Toxin level (Vintage) Must

Juice

Clarified juice

At first racking

Bottled wine

Wine after 14 mth

(a)

Proportion of OA relative to initial concentration in grapes

100% 80% 60% 40% 20% 0% Intermed (2002)

High (2002)

Intermed (2003)

High (2003)

Toxin level (Vintage) Must

After pressing

At first racking

Bottled wine

Wine after 14 mth

(b)

Figure 8. Reduction in ochratoxin A during a. white and b. red wine production. The white varieties used were Chardonnay (Chard) and Semillon (Sem) and the red variety was Shiraz. Berries which had been crushed and destemmed were termed “must”, and for this study, were deemed to contain the total ochratoxin A initially present. Only wines from 2002 vintage had been stored for 14 months at the time of analysis, hence these results are absent for the 2003 vintage.

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storage of grape juice (Roset, 2003). In most white and red ferments, racking (decanting wine after fermentation) also reduced OA content. These reductions may be due to loose interactions between OA and the solids settling out of solution. The mean OA concentration of white and red wines, respectively, at bottling was 4% and 13% of the initial concentration in grapes. Slight reductions in OA were observed in red and white wines after 14 months of storage. These trends in reduction in OA were apparently unaffected by the initial concentration of OA in the grapes, which varied in the two vintages. Initial OA concentrations fell within the range 2-66 ng/g for white grapes and 2-114 ng/g for red grapes. The reduction in OA concentration at pressing was similar for both white and red wines (approximately 70%), however, at bottling, red wines retained three fold more of the initial OA concentration than white wines. This difference may be inherent to the vinification process. In white vinification, the juice after pressing does not contain alcohol, and OA may readily bind to proteins and other solids in the juice, to be removed during clarification. In red vinification, the wine after pressing contains alcohol and hence OA present is less bound to solids and more soluble in the liquid phase. Thus racking of red wines after fermentation does not result in the same reduction in OA as clarification of white juice. Also, during fermentation of red must on skins, there may be increased partitioning of OA from the pulp and skins into the alcohol produced during fermentation. Fernandes et al. (2003) reported the opposite effect, with white wines retaining a greater proportion of initial OA than red wines (8-14% cf. 6%). This difference can be explained by noting that OA that has been spiked into crushed grapes may interact differently with grape solids compared with OA exuded directly in the berries from fungal hyphae. However, the importance of solid-liquid separations in the removal of OA during vinification is clear, regardless of the means of OA contamination.

3.4.

Future Directions

An understanding of the source of A. carbonarius and other members of Section Nigri in vineyards may aid in the development of management strategies to minimise dispersal from the soil to bunches. Reducing the frequency of tillage may be one strategy to minimise Aspergillus in the soil and air. The presence of A. carbonarius spores on bunches during the early stages of berry development does not necessarily lead to the development of Aspergillus rots and subsequent

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production of OA, as the spores do not survive well on the surface of green berries. Berry softening from veraison onwards appears to increase berry susceptibility to Aspergillus rots. Fungicide sprays are the primary tool for management of Aspergillus rots in maturing bunches, however, Australia has strict guidelines governing their use in the weeks before harvest to minimise chemical residues in the wine. The efficacy and timing of these sprays is under investigation, as part of an overall strategy to reduce the incidence of Aspergillus in vineyards. Studies of the fate of OA during vinification are also continuing, to increase understanding of the relative proportion of OA remaining in wine after vinification under Australian conditions. This has implications for setting acceptable limits for OA in winegrapes at harvest, and also for further processing of waste streams from vinification.

4.

ACKNOWLEDGEMENTS

This research was supported by the Australian Government and Australian grapegrowers and winemakers through their investment in the Cooperative Research Centre for Viticulture and Horticulture Australia Ltd. Support from grape growers in the Sunraysia district, Victoria, and Glen Howard of Somerset Vineyard, Pokolbin (Hunter Valley, New South Wales) is gratefully acknowledged, as is the contribution from Syngenta Crop Protection Pty Ltd. Narelle Nancarrow, Kathy Clarke and Margaret Leong are thanked for their help with the field trials. Winemaking was carried out with assistance from Mark Krstic, Glenda Kelly and Fred Hancock at the Victorian Department of Primary Industries, Mildura, Victoria, and Stephen W. White, Nai Tran-Dinh and Nick Charley at Food Science Australia. The key role of Peter Varelis, Kylie McClelland, Shane Cameron, Kathy Schneebeli (Analytical Chemistry, Food Science Australia) in development of the OA assays is most gratefully acknowledged. Advice on the statistical analyses was provided by John Reynolds (formerly Senior Biometrician, Victorian Department of Primary Industries, Attwood, Victoria).

5.

REFERENCES

Abarca, M. L., Accensi, F., Bragulat, M. R., and Cabañes, F. J., 2001, Current importance of ochratoxin A-producing Aspergillus spp., J. Food Prot. 64:903-906.

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Abarca, M. L., Accensi, F., Bragulat, M. R., Castellà, G., and Cabañes, F. J., 2003, Aspergillus carbonarius as the main source of ochratoxin A contamination in dried vine fruits from the Spanish market, J. Food Prot. 66:504-506. Abarca, M. L., Bragulat, M. R., Castellá, G., and Cabañes, F. J., 1994, Ochratoxin A production by strains of Aspergillus niger var. niger, Appl. Environ. Microbiol. 60:2650-2652. Abdel-Sater, M. A., and Saber, S. M., 1999, Mycoflora and mycotoxins of some Egyptian dried fruits, Bull. Fac. Sci., Assiut Univ. 28:91-107. Amerine, M. A., Berg, H. W., Kunkee, R. E., Ough, C. S., Singleton, V. L., and Webb, A. D., 1980, The Technology of Wine Making, AVI Publishing Company, Westport, CT, pp. 154-185. Anon., 2003, (March, 2004) EU food law news, FSA Letter, 29 July; http://www.food law.rdg.ac.uk/news/eu-03068.htm. Barbetti, M. J., 1980, Bunch rot of Rhine Riesling grapes in the lower south-west of Western Australia, Aust. J. Expt. Agric. Anim. Husb. 20:247-251. Battilani, P., Giorni, P., and Pietri, A., 2003a, Epidemiology of toxin-producing fungi and ochratoxin A occurrence in grape, Eur. J. Plant. Pathol. 109:715-722. Battilani, P., Pietri, A., Bertuzzi, T., Languasco, L., Giorni, P., and Kozakiewicz, Z., 2003b, Occurrence of ochratoxin A-producing fungi in grapes grown in Italy, J. Food Prot. 66:633-636. Baxter, D. E., Shielding, I. R., and Kelly, B., 2001, Behaviour of ochratoxin A in brewing, J. Am. Soc. Brew. Chem. 59:98-100. Bellí, N., Marín, S., Sanchis, V., and Ramos, A. J., 2002, Ochratoxin A (OTA) in wines, musts and grape juices: occurrence, regulations and methods of analysis, Food Sci. Technol. Int. 8:325-335. Cabañes, F. J., Accensi, F., Bragulat, M. R., Abarca, M. L., Castellá, G., Minguez, S., and Pons, A., 2002, What is the source of ochratoxin A in wine?, Int. J. Food. Microbiol. 79:213-215. Castegnaro, M. and Wild, C. P., 1995, IARC activities in mycotoxin research, Nat. Toxins 3:327-331. Castellari, M., Versari, A., Fabiani, A., Parpinello, G. P., and Galassi, S., 2001, Removal of ochratoxin A in red wines by means of adsorption treatments with commercial fining agents, J. Agric. Food Chem. 49:3917-3921. Da Rocha Rosa, C. A., Palacios, V., Combina, M., Fraga, M. E., De Oliveira Rekson, A., Magnoli, C. E., and Dalcero, A. M., 2002, Potential ochratoxin A producers from wine grapes in Argentina and Brazil, Food Addit. Contam. 19:408-414. Diehl, J. A., 1979, Influencia de sistemas de cultivo sobre podridoes de raizes de trigo. Summa Phytopathol. 5:134-139. Diehl, J. A., Tinline, R. D., Kochhann, R. A., Shipton, P. J., and Rovira, A. D., 1982, The effects of fallow periods on common root rot of wheat in Rio Grande do Sul, Brazil, Phytopathology 72:1297-1301. Duczek, L. J., 1981, Number and viability of conidia of Cochliobolus sativus in soil profiles in summer fallow fields in Saskatchewan, Can. J. Plant Path. 3:12-14. Dumeau, F., and Trione, D., 2000, Trattamenti e “ochratossina A” nei vini, Vignevini 9:79-81. Engelhardt, G., Ruhland, M., and Wallnöffer, P. R., 1999, Occurrence of ochratoxin A in moldy vegetables and fruits analysed after removal of rotten tissue parts, Adv. Food Sci. 21:88-92.

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European Commission, 2002, Commission regulation (EC) No 472/2002 of 12 March 2002 amending regulation (EC) No 466/2001 setting maximum levels for certain contaminants in foodstuffs, Off. J. Eur. Comm. 75:18-20, 42. Fernandes, A. Venâncio, A., Moura, F., Garrido, J., and Cerdeira, A., 2003, Fate of ochratoxin A during a vinification trial, Aspect. Appl. Biol. 68:73-80. Gupta, S. L., 1956, Occurrence of Aspergillus carbonarius (Bainier) Thom causing grape rot in India, Sci. Cult. 22:167-168. Heenan, C. N., Shaw, K. J., and Pitt, J. I., 1998, Ochratoxin A production by Aspergillus carbonarius and A. niger and detection using coconut cream agar, J. Food Mycol. 1:67-72. Hocking, A. D., Varelis, P., Pitt, J. I., Cameron, S., and Leong, S., 2003, Occurrence of ochratoxin A in Australian wine, Aust. J. Grape Wine Res. 9:72-78. Jarvis, W. R., and Traquair, J. A., 1984, Bunch rot of grapes caused by Aspergillus aculeatus, Plant Dis. 68:718-719. King, A. D., Hocking, A. D., and Pitt, J. I., 1981, The mycoflora of some Australian foods, Food Technol. Aust. 33:55-60. Klich, M. A., and Pitt, J. I., 1988, A Laboratory Guide to Common Aspergillus species and their Teleomorphs, CSIRO, Division of Food Processing, North Ryde, NSW, pp. 26-27, 48-51, 58-59. Leong, S. L., Hocking, A. D., and Pitt, J. I., 2004, Occurrence of fruit rot fungi (Aspergillus Section Nigri) on some drying varieties of irrigated grapes, Aust. J. Grape Wine Res. 10:83-88. MacDonald, S., Wilson, P., Barnes, K., Damant, A., Massey, R., Mortby, E., and Shepherd, M. J., 1999, Ochratoxin A in dried vine fruit: method development and survey, Food Addit. Contam. 16:253-260. Majerus, P., Bresch, H., and Otteneder, H., 2000, Ochratoxin in wines, fruit juices and seasonings, Arch. Lebensmittelhyg. 51:81-128. Marfenina, O. E., and Mirchink, T. G., 1989, Effect of human activity on soil microfungi, Sov. Soil Sci. 21:40-47. Markarki, P., Delpont-Binet, C., Grosso, F., and Dragacci, S., 2001, Determination of ochratoxin A in red wine and vinegar by immunoaffinity high-pressure liquid chromatography, J. Food Prot. 64:533-537. Nair, N. G., 1985, Fungi associated wtih bunch rot of grapes in the Hunter Valley, Aust. J. Agric. Res. 36:435-442. Pitt, J. I., and Hocking, A. D., 1997, Fungi and Food Spoilage, 2nd edition, Blackie Academic and Professional, London, pp. 385-388, 510-511. Roset, M., 2003, Survey on ochratoxin A in grape juice, Fruit Process. 13:167-172. Rotem, J., and Aust, H. J., 1991, The effect of ultraviolet and solar radiation and temperature on survival of fungal propagules, J. Phytopathol. 133:76-84. Sage, L., Krivobok, S., Delbos, É., Seigle-Murandi, F., and Creppy, E. E., 2002, Fungal flora and ochratoxin A production in grapes and musts from France, J. Agric. Food Chem. 50:1306-1311. Serra, R., Abrunhosa, L., Kozakiewicz, Z., and Venâncio, A., 2003, Black Aspergillus species as ochratoxin A producers in Portuguese wine grapes, Int. J. Food Microbiol. 88:63-68. Silva, A., Galli, R., Grazioli, B., and Fumi, M. D., 2003, Metodi di riduzione di residui di ocratossina A nei vini, Ind. Bevande 32:467-472. Snowdon, A. L., 1990, A Colour Atlas of Post-Harvest Diseases and Disorders of Fruit and Vegetables. I. General Introduction and Fruits, Wolfe Scientific, London, p. 256.

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Téren, J., Varga, J., Hamari, Z., Rinyu, E., and Kevei, F., 1996, Immunochemical detection of ochratoxin A in black Aspergillus strains, Mycopathologia 134: 171-176. Tjamos, S. E., Antoniou, P. P., Kazantzidou, A., Antonopoulos, D. F., Papageorgiou, I., and Tjamos, E. C., 2004, Aspergillus niger and Aspergillus carbonarius in Corinth raisin and wine-producing vineyards in Greece: population composition, ochratoxin A production and chemical control, J. Phytopathol. 152:250-255. Zimmerli, B., and Dick, R., 1996, Ochratoxin A in table wine and grape-juice: occurrence and risk assessment, Food Addit. Contam. 13:655-668.

OCHRATOXIN A PRODUCING FUNGI FROM SPANISH VINEYARDS Marta Bau, M. Rosa Bragulat, M. Lourdes Abarca, Santiago Minguez and F. Javier Cabañes*

1.

INTRODUCTION

Ochratoxin A (OA) is a nephrotoxic mycotoxin naturally occurring in a wide range of food commodities. It has been classified by IARC as a possible human renal carcinogen (group 2B) (Castegnaro and Wild, 1995) and among other toxic effects, is teratogenic, immunotoxic, genotoxic, mutagenic and carcinogenic (Creppy, 1999). Wine is considered the second major source of OA in Europe, with cereals being the primary source. Since the first report on the occurrence of OA in wine (Zimmerli and Dick, 1996) its presence in wine and grape juice have been reported in a broad variety of wines from different origins. Maximum OA levels have been established for cereals and dried vine fruits in the European Union, and it is possible that other commodities such as wine and grape juices will be regulated before the end of 2003 (Anonymous, 2002). Until recently, Aspergillus ochraceus and Penicillium verrucosum were considered the main OA-producing species. P. verrucosum is usually found in cool temperate regions and has been reported almost exclusively in cereal and cereal products while A. ochraceus is found

* Marta Bau, M. Rosa Bragulat, M. Lourdes Abarca and F. Javier Cabañes: Departament de Sanitat i d’Anatomia Animals, Universitat Autònoma de Barcelona, 08193 Bellaterra, Barcelona, Spain. Santiago Minguez: Institut Català de la Vinya i el Vi (INCAVI), Generalitat de Catalunya, Vilafranca del Penedés, Barcelona, Spain. Correspondence to: [email protected]

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sporadically in different commodities in warmer and tropical climates (Pitt and Hocking, 1997). The production of OA by species in Aspergillus section Nigri has received considerable attention since the first description of OA production by Aspergillus niger var. niger (Abarca et al., 1994) and by Aspergillus carbonarius (Horie, 1995). Recently, A. carbonarius and other black aspergilli belonging to the A. Niger aggregate have been described as a main possible sources of OA contamination in grapes (Da Rocha Rosa et al., 2002; Sage et al., 2002; Battilani et al., 2003; Magnoli et al., 2003; Serra et al., 2003), wine (Cabañes et al., 2002), and also in dried vine fruits (Heenan et al., 1998; Abarca et al., 2003). The objective of this study was to identify the ochratoxigenic mycobiota of grapes from vineyards mainly located along the Mediterranean coast of Spain.

2.

MATERIALS AND METHODS

2.1.

Samples

During the 2001 and 2002 seasons, fungi capable of producing ochratoxin A were isolated from the grapes from seven Spanish vineyards. The vineyards were located mainly along the Mediterranean coast and belonged to five winemaking regions: Barcelona (two vineyards), Tarragona (two vineyards), Valencia (one vineyard), Murcia (one vineyard) and Cádiz (one vineyard). In each vineyard, from May to October, samplings were made at four different times, coinciding with the following developmental stages of the grape: setting, one month after berry-set, veraison and harvesting. At each sampling time, 10 bunches were collected from 10 different plants located approximately along two crossing diagonals of the vineyard. Every bunch was collected in a separate paper bag and analyzed in the laboratory within 24-48 h of collection.

2.2.

Mycological study

Ten berries from each bunch were randomly selected, with five plated directly onto dichloran rose bengal chloramphenicol agar (DRBC) (Pitt and Hocking, 1997) and five onto malt extract agar (MEA) (Pitt and Hocking, 1997) supplemented with 100 ppm of chloramphenicol and 50 ppm of streptomycin. In total, 5,600 berries were

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analyzed. Plates were incubated at 25°C for 7 days. All fungi belonging to Aspergillus and Penicillium genera were isolated for identification to species level. (Raper and Fennell, 1965; Pitt, 1979; Klich and Pitt, 1988; Pitt and Hocking, 1997).

2.3.

Ability of Fungal Isolates to Produce Ochratoxin

Isolates belonging to Aspergillus spp. and Penicillium spp. were evaluated using a previously described HPLC screening method (Bragulat et al., 2001). Briefly, the isolates were grown on Czapek Yeast extract Agar (CYA) and on Yeast extract Sucrose agar (YES) (Pitt and Hocking, 1997) and incubated at 25°C for 7 days. Isolates identified as A. carbonarius were grown on CYA for 10 days at 30°C because these incubation conditions have been cited as optimal for detecting OA production in this species (Cabañes et al., 2002; Abarca et al., 2003). From each isolate, three agar plugs were removed from different points of the colony and extracted with 0.5 ml of methanol. The extracts were filtered and injected into the HPLC.

2.4.

Data analysis

Data obtained were analyzed statistically by means of one-way analysis of variance test and Student’s test. All statistical analyses were performed using SPSS software (version 10.0).

3.

RESULTS AND DISCUSSION

The occurrence of Aspergillus spp. in the 5,600 berries plated on the two culture media used are shown in Table 1. Although the number of isolates recovered on DRBC medium was higher than on MEA, the differences were not statistically significant. A total of 1,061 isolates belonging to twenty Aspergillus spp. (including Emericella spp. and Eurotium amstelodami) were identified. Isolates of A. carbonarius and A. niger aggregate constituted 88.7% of the total Aspergillus isolates (Figure 1). Aspergillus niger aggregate were isolated from 14.2% of plated berries, and A. carbonarius from 2.6%. The occurrence of the remaining Aspergillus spp. ranged from 0.02% to 0.5%. The distribution of the A. niger aggregate and A. carbonarius isolates in 2001 and 2002 seasons during the development of berries is shown in Figure 2. Although they were recovered in all the stages

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Table 1. Occurrence of Aspergillus spp. in grapes from Spanish vineyards examined during the 2001 and 2002 seasons No. (%) of positive berries Total DRBCa MEAa Species (n = 5,600) (n = 2,800) (n = 2,800) Aspergillus niger aggregate 797 (14.23) 438 359 A. carbonarius 144 (2.57) 83 61 A. ustus 26 (0.46) 15 11 A. fumigatus 19 (0.34) 10 9 A. flavus 11 (0.20) 6 5 A. tamarii 9 (0.16) 4 5 A. japonicus var. aculeatus 8 (0.14) 3 5 A. ochraceus 8 (0.14) 8 0 Emericella nidulans 6 (0.11) 4 2 A. alliaceus 5 (0.09) 4 1 A. terreus 5 (0.09) 4 1 A. wentii 5 (0.09) 3 2 A. melleus 4 (0.07) 4 0 A. flavipes 3 (0.05) 2 1 A. ostianus 3 (0.05) 3 0 A. parasiticus 3 (0.05) 2 1 Eurotium amstelodami 2 (0.04) 1 1 Emericella astellata 1 (0.02) 0 1 Emericella variecolor 1 (0.02) 0 1 A. versicolor 1 (0.02) 0 1 Total Aspergillus spp. 1061 594 467 a DRBC: dichloran rose bengal chloramphenicol agar; MEA: malt extract agar.

sampled, there was a statistically significant increase at harvesting. The number of isolates recovered in 2002 was lower than in 2001, probably due to different climatic conditions. Nevertheless in both seasons black Aspergilli showed the same tendency, with the highest levels of isolation at harvesting. A total of 165 isolates belonging to genus Penicillium were identified. The most frequent species were P. glabrum, P. brevicompactum, P. sclerotiorum, P. citrinum, P. chrysogenum and P. thomii. The occurrence of the remaining Penicillium spp. was lower than 0.12%. OA production was not detected by any of the 165 Penicillium isolates. Only one isolate of P. verrucosum was identified. This isolates was able to produce citrinin but did not produce OA. The ability of Aspergillus isolates to produce ochratoxin A is shown in Table 2. All the A. carbonarius isolates (n = 144) were able to produce OA whereas only eight isolates of A. niger aggregate (n = 797) were toxigenic. None of the A. japonicus var. aculeatus (n = 8) produced OA.

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Figure 1. Black aspergilli growing on plated berries from harvesting time. (Note their high occurrence at this sampling time).

A. carbonarius 2001

A. carbonarius 2002

A. niger aggregate 2001

A. niger aggregate 2002

250

No. of isolates

200 150 100 50 0 I

II

III

IV

Figure 2. Distribution of A. carbonarius and A. niger aggregate isolates at each developmental stages of the berries: I, berry set; II, one month after berry set; III, veraison; IV, harvest

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Table 2. Ochratoxin A production (µg/g of culture medium) by Aspergillus spp. isolated from Spanish grapes No. of positive Mean Species isolates /Total concentration Range A. carbonarius 144 / 144 24.6 0.1 – 378.5 A. niger aggregate 8 / 797 29.2 0.05 – 230.9 A. alliaceus 5/5 351.4 197.6 – 715.4 A. ochraceus 4/8 440.8 1.3 – 1026.7 A. ostianus 3/3 1273.3 245.9 – 2514.1 A. melleus 2/4 19.7 7.3 – 32.2

OA production was also detected by other Aspergillus species outside section Nigri. Four of the eight isolates of A. ochraceus and two of the four isolates of A. melleus produced OA. All isolates classified as A. ostianus (n=3) and A. alliaceus (n=5) were able to produce OA. Some of these species were able to produce OA in large quantities in pure culture, but due to their low occurrence, they are probably a relatively unimportant source of this mycotoxin in grapes. Although the possible participation of different OA producing species may occur, our results are strong evidence of the contribution of A. carbonarius to OA contamination in grapes, mainly at the final developmental stage of the berries, and consequently in wine.

4.

ACKNOWLEDGEMENTS

This research was supported by the European Union project QLK1-CT-2001-01761 (Quality of Life and Management of Living Resources Programme (QoL), Key Action 1 on Food, Nutrition and Health). The financial support of the Ministerio de Ciencia y Tecnología of the Spanish Government (AGL01-2974-C05-03) is also acknowledged.

5.

REFERENCES

Abarca, M. L., Bragulat, M. R., Castellá, G., and Cabañes, F. J., 1994, Ochratoxin A production by strains of Aspergillus niger var. niger, Appl. Environ. Microbiol. 60:2650-2652. Abarca, M. L., Accensi, F., Bragulat, M. R., Castellá. G., and Cabañes, F. J., 2003, Aspergillus carbonarius as the main source of ochratoxin A contamination in dried vine fruits from the Spanish market, J. Food Prot. 66:504-506.

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Anonymous, 2002. Commission regulation (EC) no 472/2002 of 12 March 2002 amending regulation (EC) no 466/2001 setting maximum levels for certain contaminants in foodstuffs, Off. J. Eur. Communities L75, 18-20. Battilani, P., Pietri, A., Bertuzzi, T., Languasco, L., Giorni, P., and Kozakiewicz, Z., 2003, Occurrence of ochratoxin A-producing fungi in grapes grown in Italy, J. Food Prot. 66:633-636. Bragulat, M. R., Abarca, M. L., and Cabañes, F. J., 2001, An easy screening method for fungi producing ochratoxin A in pure culture, Int J. Food Microbiol. 71: 139-144. Cabañes, F. J., Accensi, F., Bragulat, M. R., Abarca, M. L., Castellá, G., Minguez, S., and Pons, A., 2002, What is the source of ochratoxin A in wine?, Int J. Food Microbiol. 79:213-215. Castegnaro, M., and Wild, C. P., 1995, IARC activities in mycotoxin research, Natural Toxins 3:327-331. Creppy, E. E., 1999. Human ochratoxicoses. J. Toxicol. – Toxin Rev. 18:273-293. Da Rocha Rosa, C. A., Palacios, V., Combina, M., Fraga, M. E., De Oliveira Rekson, A., Magnoli, C. E., amd Dalcero, A. M., 2002, Potential ochratoxin A producers from wine grapes in Argentina and Brazil. Food Addit. Contam. 19:408-414. Heenan, C. N., Shaw, K. J., and Pitt, J. I., 1998, Ochratoxin A production by Aspergillus carbonarius and A. niger isolates and detection using coconut cream agar, J. Food Mycol. 1:67-72. Horie, Y., 1995, Productivity of ochratoxin A of Aspergillus carbonarius in Aspergillus section Nigri. Nippon Kingakukai Kaiho 36:73-76. Klich, M. A., and Pitt, J. I., 1988, A Laboratory Guide to Common Aspergillus Species and their Teleomorphs. CSIRO Division of Food Processing, North Ryde, NSW. Magnoli, C., Violante, M., Combina, M., Palacio, G., and Dalcero, A., 2003, Mycoflora and ochratoxin-producing strains of Aspergillus section Nigri in wine grapes in Argentina, Lett. Appl. Microbiol 37:179-184. Pitt, J. I., 1979, The Genus Penicillium and its Teleomorphic States Eupenicillium and Talaromyces, Academic Press, London. Pitt, J. I., and Hocking, A. D., 1997, Fungi and Food Spoilage, Blackie Academic and Professional, London. Raper, K. B., and Fennell, D. I., 1965, The Genus Aspergillus, The William and Wilkins Co., Baltimore. Sage, L., Krivobok, S., Delbos, E., Seigle-Murandi, F., and Creppy, E. E., 2002, Fungal flora and ochratoxin A production in grapes and musts from France, J. Agric. Food Chem. 50:1306-1311. Serra, R., Abrunhosa, L., Kozakiewicz, Z., and Venancio, A., 2003, Black Aspergillus species as ochratoxin A producers in Portuguese wine grapes, Int. J. Food Microbiol. 88:63-68. Zimmerli, B., and Dick, R., 1996, Ochratoxin A in table wine and grape-juice: occurrence and risk assessment, Food Addit. Contam. 13:655-668.

FUNGI PRODUCING OCHRATOXIN IN DRIED FRUITS Beatriz T. Iamanaka, Marta H. Taniwaki, E. Vicente and Hilary C. Menezes*

1.

INTRODUCTION

Ochratoxin A (OA) has been shown to be a potent nephrotoxin in animal species and has been found in agricultural products. OA is believed to be produced in nature by three main species of fungi, Aspergillus ochraceus, Aspergillus carbonarius and Penicillium verrucosum, with a minor contribution from A. niger and several species closely related to A. ochraceus (JECFA, 2001). P. verrucosum occur mainly in cool temperate climates, and is usually associated with cereals (Pitt, 1987; Pitt and Hocking, 1997). Studies carried out in Europe have reported the presence of the ochratoxigenic fungi A. ochraceus, A. niger and A. carbonarius and sometimes OA in dried fruits (MAFF, 2002). A. carbonarius and A. niger were described as sources of OA in maturing and drying grapes in Spain and Australia (Abarca et al., 1994; Heenan et al., 1998). Grape juice and wines from southern regions of Europe have been reported to contain detectable levels of OA (Zimmerli and Dick, 1996). Detectable concentrations of OA have been found in sultanas imported into the United Kingdom: a survey of 20 samples of dried fruit found more than 80% were positive for OA (MAFF, 1997; MacDonald et al., 1999). * Beatriz T. Iamanaka, Marta H. Taniwaki and E. Vicente: Food Technology Institute, ITAL C.P 139 CEP13.073-001 Campinas-SP, Brazil; Hilary C. Menezes: Food Engineering Faculty-Unicamp, Campinas-SP, Brazil. Correspondence to: [email protected]

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The aim of this work was to investigate the incidence of toxigenic fungi and OA in dried fruits from different countries of origin sold on the Brazilian market.

2.

MATERIALS AND METHODS

2.1.

Sampling

A total of 119 samples (500 g each) of dried fruits were purchased in Campinas and São Paulo markets in Brazil in 2002-2003, comprising black sultanas (24), white sultanas (19), apricots (14), figs (19), dates (22) and plums(21). The dried fruit samples originated from Turkey, Spain, Mexico, Tunisia, USA, Argentina and Chile.

2.2.

Mycological Analyses

Larger fruit (apricots, figs, dates and plums) were cut aseptically into small pieces, whereas smaller fruit (sultanas) were analysed whole. Whole fruit or fruit pieces were surface disinfected with 0.4% chlorine solution for 1 min. Fifty sultanas or fruit pieces were plated onto Dichloran 18% Glycerol agar (DG18; Pitt and Hocking, 1997). The plates were incubated at 25˚C for 5-7 days. Colonies with the appearance of A. niger, A. carbonarius and A. ochraceus were isolated onto Malt Extract agar (Pitt and Hocking, 1997) and identified according to Klich and Pitt (1988). The percentage infection of the fruit or pieces was calculated.

2.3.

Ochratoxin A Production

Ochratoxin A production from each isolate was analysed qualitatively using the agar plug technique of Filtenborg et al. (1983) or extracted with chloroform as described below. The isolates were inoculated onto Yeast Extract (0.1%) Sucrose (15%) agar and incubated at 25˚C for 7 days. For the agar plug technique, a small plug was cut from the colony using a cork borer and tested by TLC as described by Filtenborg et al. (1983). If isolates were found to be negative for OA production, the whole colony was extracted with chloroform. The whole colony from the Petri dish was placed in chloroform in a Stomacher and homogenised for 3 min, filtered and concentrated in a water bath at 60˚C to near dryness then dried under a stream of nitrogen. The residue was resuspended in chloroform and spotted onto TLC plates which were developed in toluene: ethyl acetate: formic acid

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(5:4:1) and visualized under UV light at 365nm. An OA standard (Sigma Chemicals, St Louis, USA) was used for comparison.

2.4.

Analysis of Ochratoxin A from Fruit Samples

The fruit samples were analysed for OA using the Ochratest™ HPLC Procedure for Currants and Raisins (Vicam, 1999). The method was validated for this study. Samples were extracted with a solution of methanol: sodium bicarbonate, 1% (70:30) in a blender at high speed for 1 min. The extracts were filtered onto qualitative paper and an aliquot diluted with phosphate buffered saline containing 0.01% Tween 20. This solution was filtered through a microfibre filter and an aliquot applied to an immunoaffinity column (Vicam, Watertown MA, USA) containing a monoclonal antibody specific for OA. The column was washed with phosphate buffered saline with 0.01% Tween 20, followed by purified water. The OA was eluted with HPLC methanol and the eluate was added to purified water, mixed and injected in the HPLC with a fluorescence detector. The mobile phase was acetonitrile:water:acetic acid (99:99:2) and the flow rate was 0.8 ml/min. The HPLC equipment was a Shimadzu LC-10VP system (Shimadzu, Japan) set at 333 nm excitation and 477 nm emission. The HPLC was fitted with a Shimadzu CLC G-ODS(4) (4 × 10 mm) guard column and Shimadzu Shimpack CLC-ODS (4.6 × 250 mm) column.

2.5.

Confirmation of Ochratoxin A

Ochratoxin A was confirmed by methyl ester formation (Pittet et al., 1996). Aliquots (200 µl) of both the sample and standard were evaporated under a stream of nitrogen and the residue redissolved in 300 µl of boron trifluoride-methanol complex (20% solution in methanol). The solution was heated at 80˚C for 10 min and allowed to cool to room temperature. The solution was evaporated and taken up in mobile phase (1 ml) and injected into the HPLC. A positive confirmation of identity was provided by the disappearance of the OA peak at retention time 16.4 min and the appearance of a new peak (OA methyl ester) at a retention time of 41.0 min.

2.6.

Water Activity

The water activity (aw) was measured using an Aqualab T3 device (Decagon, Pullman, WA, USA), at a constant temperature of 25˚C. The samples were analysed in triplicate.

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RESULTS AND DISCUSSION

The water activities of the samples examined are summarised in Table 1. Samples of all the fruit except plums and apricots were below 0.75 aw and microbiologically stable. Some samples of apricots and plums were higher than 0.75 aw, but contained preservatives and were also shelf stable. Table 2 shows the mean and range of percentage infection by A. niger plus A. carbonarius and A. ochraceus in dried fruits. The predominance of black Aspergilli can be explained by their black spores which possess resistance to ultraviolet light. The high sugar concentration and low water activity in dried fruits also assist the development of these fungi because they are xerophilic. Table 3 shows the results of the concentrations of ochratoxin A in dried fruits. The average level of contamination by OA in dried fruits was low except for one sample each of black sultanas and figs with more than 30 and 20 µg/kg OA and mean values of 4.73 and 4.10 µg/kg respectively. OA was detected at levels ranging from 0.13 to 5.0 µg/kg in most samples (88.2%). Although date samples were contaminated with a high level of toxigenic species of black Aspergilli

Table 1. Average and range of water activity (aw) in dried fruits aw Dried Fruits Mean Range Black sultanas 0.629 0.527–0.765 White sultanas 0.567 0.473–0.638 Dates 0.629 0.549–0.712 Plums 0.796 0.712–0.863 Apricots 0.694 0.638–0.782 Figs 0.682 0.638–0.751

Table 2. Percentage infection of dried fruits by A. niger plus A. carbonarius and A. ochraceus % Infection No. A. niger + A. carbonarius A. ochraceus Dried fruits Samples Mean Range Mean Range Black sultanas 24 22 0 – 90 0.8 0 –18 White sultanas 19 0.5 0–8 Dates 22 8.6 0 – 86 1.1 0 –24 Plums 21 8.0 0 – 60 0.5 0 –10 Apricots 14 Figs 19 4.5 0 – 38 -

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Table 3. Ochratoxin A levels in dried fruits Ochratoxin Black White A (µg/kg) sultanas sultanas Dates 30.0 1 Mean 4.73 0.52 150 ppm potassium sorbate. Fungal inhibition can be achieved by combining spices (or their phenolic compounds) and traditional antimicrobials, reducing the concentrations needed to achieve the same effect than when using only one antimicrobial agent. In previous studies, Azzous and Bullerman (1982) reported that clove was an efficient antimycotic agent against A. flavus, A. parasiticus and A. ochraceus and four strains of Penicillium, delaying mould growth by more than 21 days. These authors also observed additive and synergic effects combining 0.1% clove with 0.1-0.3% potassium sorbate, delaying mould germination time. Sebti and Tantaoui-Elaraki (1994) reported that the combination of sorbic acid (0.75 g/kg) with an aqueous cinnamon extract (20 g/kg) inhibited growth of 151 mould and yeast strains isolated from a Moroccan bakery product. In contrast, when using

Table 10. Fractional inhibitory concentration index (FICIndex) for Aspergillus flavus using binary mixtures of antimicrobials in potato dextrose agar formulated at aw 0.99, pH 3.5. Thymol Eugenol Thymol Carvacrol (ppm) (ppm) FICIndex (ppm) (ppm) FICIndex 50 500 0.958 100 250 1.083 100 400 0.917 150 200 1.042 150 300 0.875 200 150 1.000 200 200 0.833 250 100 0.958 250 100 0.792 300 100 1.083 300 100 0.917 350 50 1.042 350 100 1.042

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only one antimicrobial agent to inhibit the studied microorganisms, 2000 ppm of sorbic acid was needed. Matamoros-León et al. (1999) evaluated individual and combined effects of potassium sorbate and vanillin concentrations on the growth of Penicillium digitatum, P. glabrum and P. italicum in PDA adjusted to aw 0.98 and pH 3.5, and observed that 150 ppm potassium sorbate inhibited P. digitatum while 700 ppm were needed to inhibit P. glabrum. Using vanillin, inhibitory concentrations varied from 1100 ppm for P. digitatum and P. italicum to 1300 ppm for P. glabrum. When used in combination, minimal inhibitory concentration (MIC) isobolograms illustrated that curves deviated to the left of the additive line. Also, calculated FICIndex values varied from 0.60 to 0.84. FICIndex as well as isobolograms demonstrated synergistic effects on mould inhibition when vanillin and potassium sorbate were applied in combination (Matamoros-León et al., 1999).

3.2.

Ternary Mixtures

Several of the tested combinations of two antimicrobials exhibited synergy in an experimental system. The question arises as to whether combinations of more than two agents might show even greater synergy. However it is not easy to answer this question (Berenbaum et al., 1983). An alternative approach to improving fungal inhibition could be to combine three antimicrobials agents, especially for resistant strains which cannot be inhibited with individual or binary mixtures of antimicrobials. As already mentioned, little is known about the interaction between antifungal agents against filamentous fungi. In order to determine the potential use of ternary combinations of antifungal agents to inhibit growth of A. flavus we decided to study the interactions among selected antimicrobials (Table 4). Tables 11-16 present the growth/no growth results for the evaluated ternary mixtures, as well as FIC of each antimicrobial in the mixture and FICIndex. The experimental design used, proposed by Berenbaum et al. (1983), has been used for antibiotics and is focused on determining synergistic mixtures and establishing if they are consistently synergistic, i.e. results are not dependent on concentration. Several experiments (mixtures) proposed in the design (Table 4) and evaluated (Tables 11-16) have by definition an FICIndex equal to 1, and are included to corroborate individual and binary inhibitory effects, as well as ternary combinations in which the proportions of antimicrobials (fractions of MIC) add up to 1. Two thirds of the MIC of one agent and 1/6 of the MIC of the other two, or 1/3 of the MIC of each

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Table 11. Fractional inhibitory concentration (FIC) and FICIndex for Aspergillus flavus using ternary mixtures of potassium sorbate (KS), carvacrol and thymol in potato dextrose agar formulated at aw 0.99, pH 3.5. KS Carvacrol Thymol Growth FIC FIC FIC (ppm) (ppm) (ppm) Response KS Carvacrol Thymol FICIndex 400 0 0 NG 1.00 0.00 0.00 1.00 0 300 0 NG 0.00 1.00 0.00 1.00 0 0 400 NG 0.00 0.00 1.00 1.00 0 150 200 NG 0.00 0.50 0.50 1.00 200 0 200 NG 0.50 0.00 0.50 1.00 200 150 0 NG 0.50 0.50 0.00 1.00 132 99 132 NG 0.33 0.33 0.33 1.00 68 51 68 NG 0.17 0.17 0.17 0.50 68 51 268 NG 0.17 0.17 0.67 1.00 268 51 68 NG 0.67 0.17 0.17 1.00 68 201 68 NG 0.17 0.67 0.17 1.00 32 24 132 NG 0.08 0.08 0.33 0.50 132 24 32 G 32 99 32 NG 0.08 0.33 0.08 0.50

agent are examples of these ternary mixtures. The rest of the experiments are used to test synergy by combining 1/6 MIC of each agent or 1/3 MIC of one antimicrobial with 1/12 MIC of the other two. Berenbaum et al. (1983) indicated that if no growth is obtained in every combination tested, the ternary mixture is synergic in a consistent way. Table 12. Fractional inhibitory concentration (FIC) and FICIndex for Aspergillus flavus using ternary mixtures of potassium sorbate (KS), eugenol and thymol in potato dextrose agar formulated at aw 0.99, pH 3.5. KS Eugenol Thymol Growth FIC FIC FIC (ppm) (ppm) (ppm) Response KS Eugenol Thymol FICIndex 400 0 0 NG 1.00 0.00 0.00 1.00 0 600 0 NG 0.00 1.00 0.00 1.00 0 0 400 NG 0.00 0.00 1.00 1.00 0 300 200 NG 0.00 0.50 0.50 1.00 200 0 200 NG 0.50 0.00 0.50 1.00 200 300 0 NG 0.50 0.50 0.00 1.00 132 198 132 NG 0.33 0.33 0.33 1.00 68 102 68 NG 0.17 0.17 0.17 0.50 68 102 268 NG 0.17 0.17 0.67 1.00 268 102 68 NG 0.67 0.17 0.17 1.00 68 402 68 NG 0.17 0.67 0.17 1.00 32 48 132 NG 0.08 0.08 0.33 0.50 132 48 32 G 32 198 32 NG 0.08 0.33 0.08 0.50

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Table 13. Fractional inhibitory concentration (FIC) and FICIndex for Aspergillus flavus using ternary mixtures of potassium sorbate (KS), vanillin and thymol in potato dextrose agar formulated at aw 0.99, pH 3.5. KS Vanillin Thymol Growth FIC FIC FIC (ppm) (ppm) (ppm) Response KS Vanillin Thymol FICIndex 400 0 0 NG 1.00 0.00 0.00 1.00 0 1300 0 NG 0.00 1.00 0.00 1.00 0 0 400 NG 0.00 0.00 1.00 1.00 0 650 200 NG 0.00 0.50 0.50 1.00 200 0 200 NG 0.50 0.00 0.50 1.00 200 650 0 NG 0.50 0.50 0.00 1.00 132 429 132 NG 0.33 0.33 0.33 1.00 68 221 68 G 68 221 268 G 268 221 68 NG 0.67 0.17 0.17 1.00 68 871 68 NG 0.17 0.67 0.17 1.00 32 104 132 G 132 104 32 G 32 429 32 G

Therefore, above the lowest concentration of every antimicrobial tested in the mixture, the combination will be synergistic. In ternary mixtures including potassium sorbate-thymol-carvacrol (Table 11) and potassium sorbate-thymol-eugenol (Table 12), mould growth was observed only when 1/3 MIC of potassium sorbate was Table 14. Fractional inhibitory concentration (FIC) and FICIndex for Aspergillus flavus using ternary mixtures of potassium sorbate (KS), eugenol and carvacrol in potato dextrose agar formulated at aw 0.99, pH 3.5. KS Eugenol Carvacrol Growth FIC FIC FIC (ppm) (ppm) (ppm) Response KS Eugenol Carvacrol FICIndex 400 0 0 NG 1.00 0.00 0.00 1.00 0 600 0 NG 0.00 1.00 0.00 1.00 0 0 300 NG 0.00 0.00 1.00 1.00 0 300 150 NG 0.00 0.50 0.50 1.00 200 0 150 NG 0.50 0.00 0.50 1.00 200 300 0 NG 0.50 0.50 0.00 1.00 132 198 99 NG 0.33 0.33 0.33 1.00 68 102 51 NG 0.17 0.17 0.17 0.50 68 102 201 NG 0.17 0.17 0.67 1.00 268 102 51 NG 0.67 0.17 0.17 1.00 68 402 51 NG 0.17 0.67 0.17 1.00 32 48 99 G 132 48 24 G 32 198 24 NG 0.08 0.33 0.08 0.50

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Table 15. Fractional inhibitory concentration (FIC) and FICIndex for Aspergillus flavus using ternary mixtures of potassium sorbate (KS), vanillin and carvacrol in potato dextrose agar formulated at aw 0.99, pH 3.5. KS Vanillin Carvacrol Growth FIC FIC FIC (ppm) (ppm) (ppm) Response KS Vanillin Carvacrol FICIndex 400 0 0 NG 1.00 0.00 0.00 1.00 0 1300 0 NG 0.00 1.00 0.00 1.00 0 0 300 NG 0.00 0.00 1.00 1.00 0 650 150 NG 0.00 0.50 0.50 1.00 200 0 150 NG 0.50 0.00 0.50 1.00 200 650 0 NG 0.50 0.50 0.00 1.00 132 429 99 G 68 221 51 G 68 221 201 NG 0.17 0.17 0.67 1.00 268 221 51 G 68 871 51 NG 0.17 0.67 0.17 1.00 32 104 99 G 132 104 24 G 32 429 24 G

combined with 1/12 MIC of thymol and 1/12 MIC of carvacrol or eugenol. In both ternary mixtures when growth was observed the phenolic compounds represent the lowest MIC fraction tested (1/12). Combinations that result in synergism (FIC = 0.5) include at least one phenolic in a fraction higher than 1/12 MIC, Therefore, we can Table 16. Fractional inhibitory concentration (FIC) and FICIndex for Aspergillus flavus using ternary mixtures of potassium sorbate (KS), vanillin and eugenol in potato dextrose agar formulated at aw 0.99, pH 3.5. KS Vanillin Eugenol Growth FIC FIC FIC (ppm) (ppm) (ppm) Response KS Vanillin Eugenol FICIndex 400 0 0 NG 1.00 0.00 0.00 1.00 0 1300 0 NG 0.00 1.00 0.00 1.00 0 0 600 NG 0.00 0.00 1.00 1.00 0 650 300 NG 0.00 0.50 0.50 1.00 200 0 300 NG 0.50 0.00 0.50 1.00 200 650 0 NG 0.50 0.50 0.00 1.00 132 429 198 NG 0.33 0.33 0.33 1.00 68 221 102 G 68 221 402 G 268 221 102 G 68 871 102 NG 0.17 0.67 0.17 1.00 32 104 198 G 132 104 48 G 32 429 48 G

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conclude that synergistic antifungal combinations of these agents must include: 1/12 MIC < phenolic concentrations < 1/6 MIC. Comparing binary and ternary mixture results, it can be observed that a considerable reduction of potassium sorbate concentration from 150-250 ppm to 32-68 ppm is possible. Maintaining a potassium sorbate concentration near 100 ppm, allows a considerable reduction in phenolic antimicrobial concentrations in the inhibitory ternary mixtures. Figure 4 presents a representation of FIC of potassium sorbate-thymol-eugenol inhibitory combinations that inhibit A. flavus, this 3-D isobologram as well as FICIndex illustrate the synergistic combinations. Ternary mixtures where vanillin is included (Tables 13, 15 and 16) have in common that growth was observed in those combinations where synergism was expected. However, some ternary combinations presented an additive result (FICIndex = 1). Observing binary results of those mixtures that include vanillin, synergism was anticipated in ternary mixtures but the results cannot be predicted, as can be seen in Tables 13, 15 and 16. Monzón et al. (2001) reported that individual or binary antimicrobial agents that exhibit synergistic results do not necessary generate similar outcomes in ternary antimicrobial mixtures.

1

0.8

0.6 FIC KS 0.4

0.2

0.07

0.20

0.33

0.47

0.60

0.73

0.87

1

0.07

0.20

0.47

0.73

0.33

FIC Eugenol

0.60

1.00

0.87

0

FIC Thymol

Figure 4. Fractional inhibitory concentration (FIC) isobologram for potassium sorbate (KS), eugenol and thymol combinations to inhibit Aspergillus flavus in potato dextrose agar (aw 0.99 and pH 3.5) after 30 days incubation at 25°C.

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CONCLUSIONS

Combinations of antimicrobials can be selected when identified microorganisms are resistant to inhibition and/or inactivation by legal levels of single, conventional antimicrobials. The combination may exert the desired antimicrobial activity. Also, and more frequently, antimicrobial combinations can be selected to provide broad-spectrum preservation (Alzamora et al., 2003). Relatively little work has been reported to date on the combined action of mixtures of conventional and natural antimicrobials against microorganisms, even in model systems. Moreover, little attention has been given to the study of the mechanisms underlying their toxicity and modes of resistance, particularly for microorganisms of concern in fruit products, among them moulds and yeasts. This lack of understanding of the relative contribution of factors to obtain safe, high quality foods is surprising, because the necessity for an adequate database on which to develop safe multifactorial preservation systems was pointed out some time ago (Roberts, 1989). Further work to combine natural antimicrobials with conventional ones in foods is still required. Several combinations could be exploited for mild food preservation techniques in the near future. However, their mode of action in model systems and in food matrices is still not well understood and represents a barrier to their application. Results of the study reported here, and various others in the literature, raise certain questions about the use of antimicrobial mixtures (Alzamora and López-Malo, 2002; Alzamora et al., 2003). Conversely, a better understanding of microbial ecology and the physiological response of microorganisms to individual preservation factors as well as to combinations of natural and conventional antimicrobials in different food environments will offer new opportunities and provide greater precision for a rational selection of antimicrobial combinations. The answer to these points can be established by appropriate scientific and experimental inquiry and it is part of the challenge for the future of antifungal combinations.

5.

ACKNOWLEDGMENTS

We acknowledge financial support from CONACyT -Mexico (Projects 32020-B, 33405-B and 44088), Universidad de las AméricasPuebla, and CYTED Program (Project XI.15).

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PROBABILISTIC MODELLING OF ASPERGILLUS GROWTH Enrique Palou and Aurelio López-Malo*

1.

INTRODUCTION

Filamentous fungi are of concern to the food industry as potential spoilage micro-organisms (Pitt, 1989; Samson, 1989) and mycotoxin producers (Smith and Moss, 1985). It is important to understand the growth kinetics of these fungi in the food context, in order to control product quality from formulation to storage. This is especially applicable to long shelf life products, but also in those food products where formulation ingredients could be source of fungal contamination which may cause spoilage during processing or storage. In low water activity foods, mould spoilage is controlled by controlling aw, either by drying or the addition of solutes (NaCl, sucrose, glucose or fructose). However mould growth can also occur at aw values, especially when the preservation factors inhibit bacteria and allow fungal growth (Gould, 1989; Pitt and Miscamble, 1995). Mould growth in these products depends on the pH, aw and antimicrobial agents, which are directly a function of the product formulation (Rosso and Robinson, 2001), the solutes used (ICMSF, 1980; Pitt and Hocking, 1977), and the storage temperature. Several models describing the effect of aw or solute concentration on the growth of moulds have been published (Gibson et al., 1994; Cuppers et al., 1997; Valik et al., 1999; Rosso and Robinson, 2001).

*Ingeniería Química y Alimentos, Universidad de las Américas, Puebla. Cholula 72820, Mexico. Correspondence to [email protected]

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Predictive modelling as defined by the US Advisory Committee on Microbiological Criteria for Foods is the use of mathematical expressions to describe the likely behaviour of biological agents. Mathematical modelling of microbial growth or decline (also called “predictive microbiology”) is receiving a great deal of attention because of its enormous potential within the food industry. Predictive modelling provides a fast and relatively inexpensive way to get reliable first estimates on microbial growth and survival (McMeekin et al. 1993). Predictive microbiology is gaining importance as a powerful tool in food microbiology. Predictive modelling can be used for describing behaviour of microorganisms under different conditions, as well as assisting process design and optimization for production and distribution chains, based on microbial safety and shelf-life (Alavi et al., 1999; Alzamora and López-Malo, 2002). The main driving forces for the impressive progress in microbial modelling have been: the advent of reasonably priced microcomputers that has facilitated multifactorial data analysis, and the great improvement in techniques to establish mathematical models in the area of predictive microbiology. Predictive microbiology involves the use of mathematical expressions to describe microbial behaviour. These include functions that relate microbial density to time, and growth rate to environmental conditions such as temperature, pH, aw, and presence of antimicrobial agents. Predictive models in food microbiology can be divided, according to their aim, into two main categories: kinetic models and probability models. Kinetic models that predict growth of foodborne microorganisms are effective under a wide range of conditions; however, they are less useful close to the boundary between growth and no growth. Probabilistic models are useful where the objective is to determine whether or not microbial growth can occur under specific conditions. Much of the effort spent on generating predictive microbiology databases has focused on kinetic data, in which growth rates of microorganisms are determined in the normal temperature range and in combination with aw, pH and nutrient levels that do not prevent growth of the modelled organism (Salter et al., 2000). This strategy is adequate when the desired information is the extent of growth of food spoilage organisms, or of pathogens for which some tolerance of growth is acceptable. However, in many situations it is important to ensure that microorganisms do not contaminate foods (Ross and McMeekin, 1994; Tienungoon et al., 2000). The goal of analysis using any statistical model-building technique is to find the best suited and most parsimonious, yet biologically

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reasonable, model to describe the relationship between a dependent variable and a set of independent variables. However, while kinetic models make possible the calculation of the food shelf-life or the prediction of the time span in which significant microbial growth might occur, the probabilistic models focus their attention towards deciding whether a microorganism might or might not grow. Consequently, probability modelling is particularly useful when pathogenic or mycotoxin-producing species are involved. In this case, the growth rate of a microorganism is of lesser importance than the fact that it is present, and potentially able to multiply up to infectious dose or toxic levels. Ratkowsky and Ross (1995) proposed the application in food microbiology of the logistic regression model, which enables modelling of the boundary between growth and no growth for selected microbial species when one or more growth controlling factors are used. This approach was subsequently used by Presser et al. (1998), Bolton and Frank (1999), López-Malo et al. (2000), López-Malo and Palou (2000a, b), McMeekin et al. (2000), Lanciotti et al. (2001), and Palou and López-Malo (2003). In recent years, there has been a continuing interest in the development of predictive microbiology models describing microbial responses in food. The benefits of their application in the food industry could be substantial and various, such as prediction of shelf life, or as an aid to the elaboration of minimally processed foods (Alzamora and López-Malo, 2002). Several researchers have indicated that a need exists for predictive models with advantageous mathematical characteristics such as parsimony, robustness and stability (McMeekin et al., 1992; McMeekin et al., 1993; Ratkowsky, 1993; Whiting and Call, 1993; Baranyi and Roberts, 1994; Massana and Baranyi, 2000a). These properties would decrease the error of predictions and would increase the confidence in using predictive models. Multivariate polynomials are commonly used in predictive microbiology to summarise experimental results on the effect of environmental conditions on fungal growth (López-Malo and Palou, 2000b). They allow the use of linear regression for curve fitting procedures, which results in ease of computation and well established statistical analyses. The growth of microorganisms in food can be fully described only by a combination of the two kinds of models, kinetic and probabilistic. An integrated description of the microbial response could be given by first establishing the likelihood of growth through a probability model (growth/no growth boundary model), and then predicting the growth parameters; specific rate and lag time, if growth is expected (López-Malo and Palou, 2000a, b; Massana and Baranyi, 2000b;

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Palou and López-Malo, 2003). Boundary models will then help to define the range of applicability of kinetic models and may also be important for establishing food safety regulations as highlighted by Schaffner and Labuza (1997). Boundary models can predict the most suitable combinations of factors to prevent microbial growth, thus giving a significant degree of quality and safety from spoilage or food borne disease. This was also the aim of the hurdle approach proposed by Leistner (1985). Despite their potential importance, until now there have been only a few attempts to model the growth/no growth boundary for vegetative microorganisms. In this study, selected experimental designs, i.e. central composite or factorials, and the combined effects of incubation temperature, aw, pH and concentration of antimicrobial agent (vanillin or sodium benzoate) were incorporated into laboratory media to evaluate the growth/no growth response of three important mycotoxigenic Aspergillus species, namely Aspergillus flavus, A. ochraceus and A. parasiticus.

2.

MATERIALS AND METHODS

2.1.

Microorganisms and Preparation of Inocula

Aspergillus flavus ATCC 16872, A. ochraceus ATCC 22947 and A. parasiticus ATCC 26691 were cultivated on potato dextrose agar (PDA; Merck, Mexico) slants for 10 days at 25°C and the spores harvested with 10 ml of 0.1% Tween 80 (Merck, Mexico) solution sterilized by membrane (0.45 µm) filtration. Spore suspensions were adjusted with the same solution to give a final spore concentration of 106 spores/ml and were used the same day. Depending on the experimental design, a cocktail of these three species (A. flavus, A. parasiticus and A. ochraceus) or A. flavus alone were used.

2.2.

Experimental Designs

A three level central composite design (Montgomery, 1984) was employed in a first study to assess the effects of pH, incubation temperature and vanillin concentration on mould growth response at 0.98 aw. Independent variable levels are presented in Table 1. The results of this first set of experiments were used to select levels, including aw, of each variable in a range where fungi might or might not grow. A facto-

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Table 1. Central composite design utilized to evaluate growth response of a cocktail of conidia of Aspergillus flavus, A. parasiticus and A. ochraceus in laboratory media formulated with aw 0.98, selected pH, vanillin concentration and incubated at different temperatures. Incubation Vanillin Temperature Concentration pH (°C) (ppm) 3.0 10.0 0 4.0 10.0 0 3.0 25.0 0 4.0 25.0 0 2.7 17.5 0 4.3 17.5 0 3.5 4.9 0 3.5 30.1 0 3.5 17.5 0 3.0 10.0 500 4.0 10.0 500 3.0 25.0 500 4.0 25.0 500 3.0 10.0 1000 4.0 10.0 1000 3.0 25.0 1000 4.0 25.0 1000 2.7 17.5 750 4.3 17.5 750 3.5 4.9 750 3.5 30.1 750 3.5 17.5 330 3.5 17.5 1170 3.5 17.5 750

rial design was employed to assess the effects of aw, pH, antimicrobial concentration, antimicrobial type (sodium benzoate or vanillin) and incubation temperature, on Aspergillus flavus growth response, the levels of every variable are presented in Table 2. Triplicate systems were prepared with the resulting variable combinations (Tables 1 and 2).

2.3.

Laboratory Media

Following the experimental designs, PDA systems were prepared with commercial sucrose to reach aw 0.98, 0.96 or 0.94, sterilized for 15 min at 121°C, cooled and acidified with hydrochloric acid to the desired pH. The amounts of sucrose and hydrochloric acid needed in every case had been previously determined. The sterilized and acidified agar

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Table 2. Factorial design utilized to evaluate the growth response of Aspergillus flavus in laboratory media formulated with selected water activity, pH, sodium benzoate or vanillin concentration and incubated at different temperatures. 0.98 aw 0.96 aw 0.94 aw IncuIncuIncubation bation bation Temp Conca Temp Conc Temp Conc pH (°C) (ppm) pH (°C) (ppm) pH (°C) (ppm) 3 15 0 3 15 0 3 15 0 4 15 0 4 15 0 4 15 0 5 15 0 5 15 0 5 15 0 3 25 0 3 25 0 3 25 0 4 25 0 4 25 0 4 25 0 5 25 0 5 25 0 5 25 0 3 15 100 3 15 100 3 15 100 4 15 100 4 15 100 4 15 100 5 15 100 5 15 100 5 15 100 3 25 100 3 25 100 3 25 100 4 25 100 4 25 100 4 25 100 5 25 100 5 25 100 5 25 100 3 15 200 3 15 200 3 15 200 4 15 200 4 15 200 4 15 200 5 15 200 5 15 200 5 15 200 3 25 200 3 25 200 3 25 200 4 25 200 4 25 200 4 25 200 5 25 200 5 25 200 5 25 200 . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 15 1000 3 15 1000 3 15 1000 4 15 1000 4 15 1000 4 15 1000 5 15 1000 5 15 1000 5 15 1000 3 25 1000 3 25 1000 3 25 1000 4 25 1000 4 25 1000 4 25 1000 5 25 1000 5 25 1000 5 25 1000 a Concentration of sodium benzoate or vanillin

solutions were aseptically divided and depending on the experimental design, the necessary amount of vanillin or sodium benzoate (Sigma Chemical, Co., St. Louis, MO, was added and mechanically incorporated under sterile conditions, then poured into sterile Petri dishes.

2.4.

Inoculation and Incubation

Triplicate Petri dishes of every system were centrally inoculated by pouring 2 µl of the spore suspension (≈ 2.0 × 103 spores/plate) to give

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a circular inoculum (1 mm diameter). For every tested pH and aw, growth controls without antimicrobial were prepared and inoculated as above, including one control without pH adjustment (pH = 5.5) or aw change (aw = 0.998). Three plates of each system were maintained without inoculation for aw and pH measurement. The inoculated plates and controls were incubated for 1 month at selected temperatures (Tables 1 and 2) in hermetically closed plastic containers to avoid dehydration. A sufficient headspace was left in the containers to avoid anoxic conditions. Periodically, inoculated plates were removed briefly to observe them and determine if growth had occurred and immediately re-incubated. Water activity was measured with a Decagon CX1 instrument (Decagon Devices, Inc., Pullman, WA) calibrated and operated following the procedure described by López-Malo et al. (1993). pH was determined with a Beckman pH meter model 50 (Beckman Instruments, Inc., Fullerton, CA). Measurements were made by triplicate. The pH and aw of the PDA systems without inoculation determined at the beginning and at the end of incubation demonstrated that the desired values remained constant under incubation conditions.

2.5.

Mould Growth Response

The inoculated systems were examined daily using a stereoscopic microscope (American Optical, model Forty). A diameter of approximately 2 mm was defined as a positive sign of growth (López-Malo et al., 1998) and registered as “1.” If no growth was observed during the incubation period (one month), the response was registered as “0”.

2.6.

Model Construction

A logistic regression model relates the probability of occurrence of an event, Y, conditional on a vector, x, of explanatory variables (Hosmer and Lemeshow, 1989). The quantity p (x) = E (Y⎪x) represents the conditional mean of Y given x when the logistic distribution is used. The specific model of the logistic regression is as follows: p (x) = [exp (Σ bi xi)] / [1+ exp (Σ bi xi)]

(1)

The logit transformation of p (x) is defined as: logit (p) = g (x) = ln {[p (x) ] / [1-p (x)]} = Σ bi xi

(2)

For our particular case, aw, pH, incubation temperature (T), antimicrobial type (A), antimicrobial concentration (C) and their

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interactions are the independent variables and the outcome or dependent variable is mould response. In order to fit the logistic model the following equations were selected: Central composite design – Aspergillus flavus, A. ochraceus and A. parasiticus growth response g(x)=β0 +β1pH+β2pH2 +β3T +β4T2 + β5C +β6C2 + β7pHT+ β8pHC+β9TC +β10pHTC

(3)

Factorial design -Aspergillus flavus growth response g(x)=β0 + β1T +β2 aw +β3 pH+β4 C +β5 A +β6 Taw + β7 TpH+ β8 TC +β9 TA +β10 aw pH+β11 aw C +β12 aw A +β13 pHC+ β14 pHA+β15 CA+ β16 Taw pH+ β17 TpHC+ β18 TCA +β19 Taw C +β20 TpHA+β21 aw pHC+ β22 awCA+β23 pHCA+ β24 Taw pHC+ β25 Taw pHA+ β26 aw pHCA+β27 Taw pHCA (4)

where the coefficients (βi) are the parameters to be estimated by fitting the models to our experimental data. If an independent variable is discrete, then it is inappropriate to include it in the model as if it was interval scaled. In this situation, the method of choice is to use a collection of design or dummy variables (Hosmer and Lemeshow, 1989). For antimicrobial type (A) which is a discrete variable the codification we used was as follows: “1” for vanillin and “0” for sodium benzoate. Logistic regression was performed with the logistic subroutine in SPSS 10.0 (SPSS Inc., Chicago, IL). A forward stepwise selection procedure was performed to fit the logistic regression equation. The significance of the coefficients was evaluated and were eliminated from the model if the probability of being zero was greater than 0.1. After fitting the logistic regression equation, predictions of the growth/no growth interface were made at probability levels of 0.50, 0.10 and 0.05, by substituting the value of logit (p) in the model and finding the value of one independent variable maintaining fixed the other independent variables. Also probability of growth was calculated using the logistic equation for the evaluated conditions.

3.

RESULTS AND DISCUSSION

For every tested pH and aw, controls prepared without antimicrobials produced growth when the incubation temperature was 15°C or

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higher, but at temperatures of 10°C or lower no growth was observed after one month of incubation (Table 3). Growth/no growth results for the conditions of central composite design (Table 1) and factorial design (Table 2) are summarized in Tables 3 and 4, respectively. Results demonstrate that mould inhibition (no observable growth) can be obtained with several pH, incubation temperature and vanillin concentration combinations (Table 3) or with combinations of aw, pH, incubation temperature and antimicrobial (sodium benzoate or vanillin) concentration (Table 4). In most cases once a replicate from a combination produced growth eventually all the others also grew. Therefore, the probabilities observed were, with some exceptions, either close to 1 or 0. This observation agrees with Ratkowsky et al. (1991), they reported higher variance of the microbial response under more stressful conditions. Some synergistic combinations can be

Table 3. Response (growth=1, no growth=0) of an Aspergillus flavus A. parasiticus and A. ochraceus cocktail inoculated in potato dextose agar formulated at aw 0.98, selected pH values and different concentrations of vanillin incubated at different temperatures. Incubation Vanillin Mould Cocktail pH Temperature (°C) Concentration (ppm) Growth Response 2.7 17.5 0 1 2.7 17.5 750 0 3.0 10.0 0 0 3.0 10.0 500 0 3.0 10.0 1000 0 3.0 25.0 0 1 3.0 25.0 500 1 3.0 25.0 1000 0 3.5 4.9 0 0 3.5 4.9 750 0 3.5 17.5 0 1 3.5 17.5 330 1 3.5 17.5 750 1 3.5 17.5 1170 0 3.5 30.1 0 1 3.5 30.1 750 1 4.0 10.0 0 0 4.0 10.0 500 0 4.0 10.0 1000 0 4.0 25.0 0 1 4.0 25.0 500 1 4.0 25.0 1000 1 4.3 17.5 0 1 4.3 17.5 750 1

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Table 4. Response (growth=1, no growth=0) of Aspergillus flavus inoculated in potato dextrose agar formulated with selected aw, pH values and different concentrations of sodium benzoate or vanillin incubated at 25 or 15°C. 0.94 aw 0.96 aw 0.98 aw Temp Conc.a Sodium Sodium Sodium (°C) pH (ppm) Benzoate Vanillin Benzoate Vanillin Benzoate Vanillin 25 3 0 1 1 1 1 1 1 200 1 1 1 1 1 1 400 0 1 0 1 1 1 600 0 1 0 1 0 1 800 0 1 0 1 0 1 1000 0 0 0 0 0 0 4 0 1 1 1 1 1 1 200 1 1 1 1 1 1 400 1 1 0 1 1 1 600 0 1 0 1 1 1 800 0 1 0 1 0 1 1000 0 1 0 1 0 1 5 0 1 1 1 1 1 1 200 1 1 1 1 1 1 400 1 1 1 1 1 1 600 1 1 1 1 1 1 800 1 1 1 1 1 1 1000 1 0 1 1 1 1 15 3 0 1 1 1 1 1 1 200 0 1 0 1 1 1 400 0 0 0 0 0 0 600 0 0 0 0 0 0 800 0 0 0 0 0 0 1000 0 0 0 0 0 0 4 0 1 1 1 1 1 1 200 0 1 1 1 1 1 400 0 0 0 0 0 1 600 0 0 0 0 0 0 800 0 0 0 0 0 0 1000 0 0 0 0 0 0 5 0 1 1 1 1 1 1 200 0 1 1 1 1 1 400 0 0 0 0 0 1 600 0 0 0 0 0 1 800 0 0 0 0 0 0 1000 0 0 0 0 0 0 a Concentration of antimicrobial compound (sodium benzoate or vanillin)

detected in Tables 3 and 4 where fungal growth was inhibited at relatively high pH values or incubation temperatures when combined with reduced aw and selected antimicrobial concentrations. This is the principle of the multifactorial preservation (or hurdle technology)

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approach: the search for those “minimal” combinations of factors that inhibit microbial growth. Fitting Eqs. (3) and (4) to the growth/no growth data by logistic regression and eliminating non-significant (p>0.10) terms resulted in the reduced models presented in Table 5. Variables included in the models were statistically significant (p>0.005). The goodness of fit of every model was tested by log likelihood ratio and Chi-square tests, both being significant, which indicates that models are useful to predict the outcome variable (growth or no growth). The models’ goodness of fit was also evaluated comparing predicted values and experimental observations. A predicted probability of growth (cut value) ≥ 0.50 was considered as a growth prediction. Using this criterion, the overall correct observation was 99.2% for the central composite design; with only 3 misclassified predictions from a total of 171 observations. In only one of these three disagreements was growth predicted when no growth was observed, and in the other two cases the model predicted no growth when growth was observed. For the factorial experimental design, the overall correct observation was

Table 5. Reduced logistic model coefficients utilized to predict mould growth/nogrowth for the evaluated experimental designs. Factorial design Central composite design Coefficient Term Estimate Coefficient Term Estimate β0 Constant −9876.684 β0 Constant −201.673 β1 T 744.823 β5 C 1070.559 β2 aw 9985.567 β7 pH*T 4.624 β3 pH 2182.488 β9 T*C −109.451 β5 A −7002.668 β6 T*aw −753.751 β7 T*pH −156.855 β8 T*C −0.336 β9 T*A −19.818 β10 aw*pH −2201.049 β12 aw*A 7619.495 β13 pH*C −1.523 β15 C*A −1.274 β16 T*aw*pH 158.516 β17 T*pH*C 0.126 β18 T*C*A 0.051 β19 T*aw*C 0.340 β20 T*pH*A 63.797 β21 aw*pH*C 1.541 β24 T*aw*pH*C −0.128 β25 T*aw*pH*A −64.832

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98.0%, with only 13 misclassified predictions from a total of 648 observations. In ten of these 13 discrepancies, growth was predicted when no growth was observed, and in the other three cases the model predicted no growth when growth occurred. Probabilistic microbial models based on logistic regression have been reported for Shigella flexneri (Ratkowsky and Ross, 1995), Escherichia coli (Presser et al., 1998), Saccharomyces cerevisiae (López-Malo et al., 2000), Listeria monocytogenes (Bolton and Frank, 1999) and Zygosaccharomyces bailii (Cole et al., 1987; López-Malo and Palou, 2000a, b). These reports illustrate the flexibility of logistic regression in constructing the model, taking into account square root type kinetic models (Ratkowsky and Ross, 1995; Presser et al., 1998) or polynomial type models (Cole et al., 1987; Bolton and Frank, 1999; López-Malo et al., 2000; López-Malo and Palou, 2000a, b). However, in every case growth/no growth observations were recorded at fixed storage times, which limited probabilistic models to predict microbial response during that specific period of time. Polynomial type models do not contribute to the understanding of the mechanism involved in microbial growth inhibition. However, they are useful to determine independent variable effects and their interactions. As reported for Z. bailii, the probability of growth depends on individual effects of pH, °Brix, sorbic acid, benzoic acid and sulfite concentrations as well as upon complex interactions among preservation factors (Cole et al., 1987). For Saccharomyces cerevisiae, polynomial probabilistic models predicted the boundary between growth/no growth interface of as a function of aw, pH and potassium sorbate concentration (López-Malo et al., 2000). The predicted probabilities of growth for a cocktail of Aspergillus flavus, A. ochraceus and A. parasiticus are given in Figures 1 and 2, and for Aspergillus flavus after one month of incubation in Figures 3 and 4. pH reduction gradually increased the number of combinations of incubation temperature and antimicrobial concentration with probabilities >0.05 for inhibition of mould growth. An important shift of probability of growth curves is also observed with increasing vanillin concentration (Figures 1 and 2). From Figures 3 and 4, the magnitude of the shift in the predicted boundary depends on the combination of temperature, antimicrobial type and water activity considered. The transition from “likely to grow” conditions (p > 0.90, or > 90% likelihood of growth) to “unlikely to grow” conditions (p < 0.10, or < 10% likelihood of growth), as predicted from the fitted models, was abrupt as can be seen graphically for combinations of pH and temperatures (Figures 1 and 2) as well as for pH and antimicrobial

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1.0 0.9 0.8 0.7 0.6 p

0.5 0.4 0.3 30 25

0.2 20

0.1 15

1 4.

9 3.

7

5

3 3.

3.

5

pH

3.

1 3.

9 2.

2.

10

7

0.0

Temperature (⬚C)

Figure 1. Aspergillus flavus, A. ochraceus and A. parasiticus cocktail probability of growth (p) in potato dextrose agar formulated at aw 0.98, selected pH values and 750 ppm vanillin after one month of incubation at different temperatures.

concentration (Figures 3 and 4). The abruptness of the transition between growth or no growth conditions influenced by pH can be as little as 0.1 to 0.2 pH units, which is close to the limit of reproducibility for pH measurements. For temperature the transition is much less abrupt, occurring over increments of temperature that exceed that of measurement or experimental error. For 648 factor combinations for Aspergillus flavus, only 15 of these combinations gave a response different from “all grew” or “none grew”. Thus, the experimental data showed an abrupt transition between growth and no growth. This abruptness does indicate a microbiological reality in which small changes in environmental factors within an experiment may have a strong influence on the position of the interface (Tienungoon et al., 2000; Masana and Baranyi, 2000a, b). An important feature of the generated probabilistic models is that the level of probability can be set, depending on the level of stringency required, to calculate critical values of selected variables. To illustrate, three sets of model predictions using factorial design results were compared, p = 0.50, p = 0.10 and p = 0.05. More stringent values

1.0 0.9 0.8 0.7 0.6 p 0.5 0.4 0.3 30 25

0.2

Temperature (ⴗC)

20

0.1

15

4.

1

9 3.

7 3.

5

3

3.

1

pH

5

3.

3.

9 2.

2.

10

7

0.0

Figure 2. Aspergillus flavus, A. ochraceus and A. parasiticus cocktail probability of growth (p) in potato dextrose agar formulated with aw 0.98, selected pH values and 1000 ppm vanillin concentration after one month of incubation at different temperatures.

1 0.9 0.8 0.7 0.6 p 0.5 0.4 0.3 0.2 0.1

10 00

90 0

80 0

70 0

0

4 0 3.

concentration (ppm)

3.

60

50 0

40 0

30 0

20 0

10 0

0 5. 6 4. 2 4. 8 3.

0

0

pH

Figure 3. Aspergillus flavus probability of growth (p) in potato dextrose agar formulated with selected pH, sodium benzoate concentration (ppm) and aw 0.94 after one month of incubation at 25°C.

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1 0.9 0.8 0.7 0.6 p

0.5 0.4 0.3 0.2 0.1

0

0

0 0

0 0 00 10

0

90

pH

3.

80

4

70

0

0 60

50

40

30

0

8

3. 3.

concentration (ppm)

2

4.

20

10

4.

0 5. 6

0

0

Figure 4. Aspergillus flavus probability of growth (p) in potato dextrose agar formulated with selected pH, vanillin concentration (ppm) and aw 0.98 after one month of incubation at 15°C.

(p = 0.01 or 0.001) may be necessary in some instances. Critical antimicrobial concentration predictions (Tables 6 and 7) made at p = 0.50 (50:50 chance that A. flavus will grow) represent a relatively conservative series of estimates, with the predicted response on the boundary being no better than a coin toss. Model predictions were made more stringent by making p = 0.10 or 0.05 (10 or 5% chance of a false prediction) which causes a shift in the predicted critical antimicrobial concentration or, in other words, in the growth/no growth boundary to a lower temperature, pH and water activity. As aw decreases from 0.98 (Table 6) to 0.94 (Table 7), the critical concentration of sodium benzoate or vanillin decreases. Also as incubation temperature increases higher antimicrobial concentrations are needed to achieve no growth results after one month of incubation.

4.

CONCLUSIONS

Traditional preservation and storage procedures to produce safe and stable food product are generally based on microbial control. If

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Table 6. Critical antimicrobial concentrations (ppm) for selected probabilities (p) to inhibit Aspergillus flavus growth in laboratory media formulated at aw 0.98 and selected pH values during one month of incubation at various temperatures. Incubation Temperature (°C) p pH 15.0 17.5 20.0 22.5 25.0

Sodium Benzoate 0.05 3.0 3.5 4.0 4.5 5.0

353 360 368 377 385

357 365 373 381 389

364 372 381 390 399

384 393 403 415 427

566 656 860 > 1000 > 1000

3.0 3.5 4.0 4.5 5.0

351 359 367 375 383

355 363 371 379 387

361 370 378 387 396

379 388 398 409 422

542 623 806 > 1000 > 1000

3.0 3.5 4.0 4.5 5.0

347 355 363 371 379

349 357 365 374 382

354 362 370 379 388

365 374 383 393 405

471 525 646 > 1000 > 1000

3.0 3.5 4.0 4.5 5.0

392 473 569 685 829

532 595 679 799 984

682 741 834 1004 > 1000

842 921 > 1000 > 1000 > 1000

> 1000 > 1000 > 1000 > 1000 > 1000

3.0 3.5 4.0 4.5 5.0

369 448 542 655 795

509 567 647 760 934

658 711 796 950 > 1000

817 888 > 1000 > 1000 > 1000

988 > 1000 > 1000 > 1000 > 1000

3.0 3.5 4.0 4.5 5.0

302 375 462 567 696

439 488 553 646 790

586 624 683 790 > 1000

743 790 880 > 1000 > 1000

911 998 > 1000 > 1000 > 1000

0.10

0.50

Vanillin 0.05

0.10

0.50

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Table 7. Critical antimicrobial concentrations (ppm) for selected probabilities (p) to inhibit Aspergillus flavus growth in laboratory media formulated at aw 0.94 and selected pH values during one month of incubation at various temperatures. Incubation Temperature (°C) p pH 15.0 17.5 20.0 22.5 25.0

Sodium Benzoate 0.05 3.0 3.5 4.0 4.5 5.0

50 79 108 139 170

68 94 122 151 183

98 120 146 175 209

152 173 200 236 287

291 341 436 701 > 1000

3.0 3.5 4.0 4.5 5.0

49 78 107 137 169

67 93 120 150 182

96 119 144 173 207

150 171 197 233 282

287 335 428 685 > 1000

3.0 3.5 4.0 4.5 5.0

46 75 104 134 165

63 89 116 146 177

91 113 138 167 201

143 163 188 222 269

274 318 403 638 > 1000

3.0 3.5 4.0 4.5 5.0

232 233 235 236 238

410 383 349 307 251

599 558 502 417 275

799 767 714 614 345

> 1000 > 1000 > 1000 > 1000 > 1000

3.0 3.5 4.0 4.5 5.0

228 229 230 232 233

406 379 344 301 244

595 554 496 410 265

795 761 707 604 326

1007 > 1000 > 1000 > 1000 > 1000

3.0 3.5 4.0 4.5 5.0

217 217 217 218 219

394 365 330 284 225

582 539 479 388 237

782 746 687 574 271

994 995 997 1001 > 1000

0.10

0.50

Vanillin 0.05

0.10

0.50

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the microbial hazard or spoilage cannot be totally eliminated from the food, microbial growth and toxin production must be inhibited. Microbial growth can be inhibited by combining intrinsic food characteristics with extrinsic storage and packaging conditions. Accurate quantitative data about the effects of combined factors on growth or survival of selected microorganisms are needed. Predictive models can provide decision support tools for the food industry. In many cases models are empirical, interpreting only the response of the microorganism without understanding the mechanism of the response. However, if the models are used properly, predictive probabilistic models are helpful tools for evaluating microbial responses which can in turn identify potential problems for a product, process or storage conditions. Logistic regression is a useful tool for modelling the boundary between growth and no growth. The probabilistic microbial modelling approach can provide a practical means of evaluating the combined effects of food formulation, processing and storage conditions.

5.

ACKNOWLEDGMENTS

We acknowledge financial support from CONACyT -Mexico (Projects 32405-B and 44088), Universidad de las Américas-Puebla, and CYTED Program (Project XI.15).

6.

REFERENCES

Alavi, S. H., Puri, V. M., Knabel, S. J., Mohtar, R. H., and Whiting, R. C., 1999, Development and validation of a dynamic growth model for Listeria monocytogenes in fluid whole milk, J. Food Prot. 62:170-176. Alzamora, S.M., and López-Malo, A., 2002, Microbial behavior modeling as a tool in the design and control of minimally processed foods, in: Engineering and Food for the 21st Century, J. Welti-Chanes, G. V. Barbosa-Canovas, and J. M. Aguilera, eds, CRC Press, Boca Raton, FL, pp. 631-650. Baranyi, J., and Roberts, T. A., 1994, A dynamic approach to predicting bacterial growth in food, Int. J. Food Microbiol. 23:277-294. Bolton, L. F., and Frank, J. F., 1999, Defining the growth/no growth interface for Listeria monocytogenes in Mexican-style cheese based on salt, pH and moisture content, J. Food Prot. 62:601-609. Cole, M. B., Franklin, J. G., and Keenan, M. H. J., 1987, Probability of growth of the spoilage yeast Zygosaccharomyces bailii in a model fruit drink system, Food Microbiol. 4:115-119.

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Cuppers, H. G. M., Oomes, S., and Brul, S., 1997, A model for the combined effects of temperature and salt concentration on growth rate of food spoilage moulds, Appl. Environ. Microbiol. 63:3764-3769. Gibson, A. M., Baranyi, J., Pitt, J. I., Eyles, M. J., and Roberts, T. A., 1994, Predicting fungal growth: effect of water activity on Aspergillus flavus and related species, Int. J. Food Microbiol. 23:419-431. Gould, G. W., 1989, Mechanisms of Action of Food Preservation Procedures, Elsevier, London. Hosmer, D. W., and Lemeshow, S., 1989, Applied Logistic Regression, John Wiley and Sons, New York, p. 307. ICMSF (International Commission on the Microbiological Specifications for Foods), 1980, Microbial Ecology of Foods, Vol. 1. Factors Affecting Life and Death of Microorganisms. ICMSF, Academic Press, New York. Lanciotti, R., Sinigaglia, M., Gardini, F., Vannini, L., and Guerzoni, M. E., 2001, Growth/no growth interfaces of Bacillus cereus, Staphylococcus aureus and Salmonella enteritidis in model systems based on water activity, pH, temperature and ethanol concentration, Food Microbiol. 18:659-668. Leistner, L., 1985, Hurdle technology applied to meat products of shelf stable and intermediate moisture food types, in: Properties of Water in Foods in Relation to Quality and Stability, D. Simatos, and J. L. Multon, ed., Martinus Nihof Publishers, Dordrecht, The Netherlands, pp. 309-329. López-Malo, A., and Palou E., 2000a, Growth/no growth interface of Zygosaccharomyces bailii as a function of temperature, water activity, pH, potassium sorbate and sodium benzoate concentration, Presented at Predictive Modeling in Foods, Leuven, Belgium, September 12-15. López-Malo, A., and Palou E., 2000b, Modeling the growth/no growth interface of Zygosaccharomyces bailii in mango puree, J. Food Sci. 65:516-520. López-Malo, A., Alzamora, S. M., Argaiz, A., 1998, Vanillin and pH synergistic effects on mold growth, J. Food Sci. 63:143-146. López-Malo, A., Guerrero, S., and Alzamora, S. M., 2000, Probabilistic modeling of Saccharomyces cerevisiae inhibition under the effects of water activity, pH and potassium sorbate, J. Food Prot. 63:91-95. López-Malo, A., Palou, E., and Argaiz, A., 1993, Medición de la actividad de agua con un equipo electrónico basado en el punto de rocío, Información Tecnológica. 4(6): 33-37. Masana, M. O., and Baranyi, J., 2000a, Adding new factors to predictive models: the effect on the risk of extrapolation, Food Microbiol. 17:367-374. Masana, M. O., and Baranyi, J., 2000b, Growth/no growth interface of Brochothrix thermosphacta as a function of pH and water activity, Food Microbiol. 17:485-493. McMeekin, T. A., Olley, J., Ross, T., and Ratkowsky, D. A., 1993, Predictive Microbiology: Theory and Application, Research Studies Press, Tauton, UK. McMeekin, T. A., Presser, K., Ratkowsky, D. A., Ross, T., Salter, M., and Tienungoon, S., 2000, Quantifying the hurdle concept by modelling the growth/no growth interface. A review, Int. J. Food Microbiol. 55:93-98. McMeekin, T. A., Ross, T., and Olley, J., 1992, Application of predictive microbiology to assure the quality and safety of fish and fish products, Int. J. Food Microbiol. 15:13-32. Montgomery, D. C., 1984, Design and Analysis of Experiments, John Wiley and Sons, New York.

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Palou, E., and López-Malo, A., 2004, Growth/no-growth interface modeling and emerging technologies, in: Novel Food Processing Technologies, G. V. BarbosaCanovas, M. S. Tapia, and P. Cano, eds, Marcel Dekker, New York, pp. 629-651. Pitt, J. I., 1989, Food mycology – an emerging discipline, Soc. Appl. Bacteriol. Symp. Suppl. 1989:1S-9S. Pitt, J. I., and Hocking, A. D., 1977, Influence of solute and hydrogen ion concentration on the water relations of some xerophilic fungi, J. Gen. Microbiol. 101:35-40. Pitt, J. I., and Miscamble, B. F., 1995, Water relations of Aspergillus flavus and closely related species, J. Food Prot. 58:86-90. Presser, K. A., Ross, T., and Ratkowsky, D. A., 1998, Modelling of the growth limits (growth/no-growth) of Escherichia coli as a function of temperature, pH, lactic acid concentration, and water activity, Appl. Environ. Microbiol. 64:1773-1779. Ratkowsky, D. A., and Ross, T., 1995, Modelling the bacterial growth/no-growth interface, Lett. Appl. Microbiol. 20:29-33. Ratkowsky, D. A., 1993, Principles of modelling, J. Indust. Microbiol. 12:195-199. Ratkowsky, D. A., Ross, T., McMeekin, T. A., and Olley, J., 1991, Comparison of Arrhenius-type and Belehradek-type models for prediction of bacterial growth in foods, J. Appl. Bacteriol. 71:452-459. Ross, T., and McMeekin, T. A., 1994, Predictive microbiology, Int. J. Food Microbiol. 23:241-264. Rosso, L., and Robinson, T. P., 2001, Cardinal model to describe the effect of water activity on the growth of moulds, Int. J. Food Microbiol. 63:265-273. Salter, M. A., Ratkowsky, D. A., Ross, T., and McMeekin, T. A., 2000, Modelling the combined temperature and salt (NaCl) limits for growth of a pathogenic Escherichia coli strain using nonlinear logistic regression, Int. J. Food Microbiol. 61:159-167 Samson, R. A., 1989, Filamentous fungi in food and feed, Soc. Appl. Bacteriol. Symp. Suppl. 1989:27S-35S. Schaffner, D. W., and Labuza, T. P., 1997, Predictive microbiology: where are we, and where are we going?, Food Technol. 51:95-99. Smith, J. E., and Moss, M .O., 1985, Mycotoxins, Formation, Analysis and Significance, John Wiley and Sons, Chichester, UK. Tienungoon, S., Ratkowsky, D. A., McMeekin, T. A., and Ross, T., 2000, Growth limits of Listeria monocytogenes as a function of temperature, pH, NaCl, and lactic acid, Appl. Environ. Microbiol. 11:4979-4987. Valik, L., Baranyi, J., and Gorner, F., 1999, Predicting fungal growth: the effect of water activity on Penicillium roqueforti, Int. J. Food Microbiol. 47:141-146. Whiting, R. C., and Call, J. E., 1993, Time of growth model for proteolytic Clostridium botulinum, Food Microbiol. 10:295-301.

ANTIFUNGAL ACTIVITY OF SOURDOUGH BREAD CULTURES Lloyd B. Bullerman, Marketa Giesova, Yousef Hassan, Dwayne Deibert and Dojin Ryu*

1.

INTRODUCTION

Many strategies have been studied for control of mould growth and reduction in mycotoxin production in foods. The most effective strategy for controlling the presence of mycotoxins in foods is prevention of growth of the mycotoxin-producing fungi in foods and field crops in the first place. Mycotoxin contamination may occur prior to harvest of crops and is often the dominant reason for the occurrence of mycotoxins in foods and feeds. However, fungal growth on stored foods and commodities is also a serious and continuing problem. In recent years increased public concern over chemical food additives and fungicides in foods has prompted searches for safe naturally occurring biological agents with antifungal potential. One source of such compounds are the lactic acid bacteria. While only a relatively limited number of studies have reported the inhibitory effects of lactic acid bacteria on fungal growth and mycotoxin production, it is generally believed that it is safe for humans to consume lactic acid bacteria and has been known for many years that lactic acid bacteria may positively influence the gastrointestinal tract

* Lloyd B. Bullerman, Yousef Hassan, Dwayne Deibert and Dojin Ryu, Department of Food Science and Technology, University of Nebraska, Lincoln, NE 68583-0919, USA. Marketa Giesova, Department of Dairy and Fat Technology, Institute of Chemical Technology, Prague, Czech Republic. Correspondence to [email protected]

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of humans and other mammals (Sandine, 1996). Lactic acid bacteria have been used to ferment foods for centuries, which suggests the nontoxic nature of metabolites produced by these bacteria (Garver and Muriana, 1993; Klaenhammer, 1998). Indigenous lactic acid bacteria are commonly found in retail foods, which suggests that the public consumes viable lactic acid bacteria in many ready-to-eat products (Garver and Muriana, 1993). Thus the metabolic activity of lactic acid bacteria that may contribute in a number of ways to the control of bacterial pathogens and might also have applications for preventing fungal growth (Gourama and Bullerman, 1995; Holzapfel et al., 1995; Klaenhammer, 1998). The potential of lactic acid bacteria for use as biological control agents of moulds in barley and thus improve the quality and safety of malt during the malting process has been studied in Finland (Haikara et al., 1993; Haikara and Laitila, 1995). The latter authors found a group of lactic acid bacteria with antagonistic activities against Fusarium. Lactobacillus planarum and Pediococcus pentosaceum inhibited Fusarium avenaceum obtained from barley kernels. The preliminary characterization of these starter cultures revealed new types of antimicrobial substances with low molecular mass and features not previously reported for lactic acid bacteria microbiocides (Haikara and Niku-Paavola, 1993; Niku-Paavola et al., 1999). Studies of Lactobacillus strains that possess antifungal properties carried out in the Czech Republic showed that Lactobacillus rhamnosus VT1 exhibited strong antifungal properties (Stiles et al., 1999; Plockova et al., 2000, 2001). Further research has shown that L. rhamnosus is capable of inhibiting the growth of many spoilage and toxigenic fungi including species in the genera Aspergillus, Penicillium and Fusarium (Stiles et al., 2002). The use of sourdough bread cultures has been reported to increase the shelf life of baked goods by delaying mould growth due to the presence of lactic acid bacteria (Gobbetti, 1998). This activity has been attributed to the presence of organic acids, particularly lactic and acetic acids (Spicher, 1983; Rocken, 1996). Further studies have shown that although acetic acid contributes to the antifungal activity of lactic acid bacteria, other bacterial metabolites also have antifungal activity and may contribute to the inhibition of mould growth (Corsetti et al., 1998; Gourama, 1997; Niku-Paavola et al., 1999). Strains of Lactobacillus plantarum in particular seem to possess strong antifungal activity (Gobetti et al., 1994a,b; Karunaratne et al., 1990). Lavermicocca et al. (2000) found that L. plantarum from sourdough was fungicidal to F. graminearum in wheat flour hydrolysate and

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reported the purification and characterization of novel antifungal compounds from this strain. The compounds that they reported to have strong antifungal activity were phenyllactic acid and 4-hydroxyphenyllactic acid. Two strains of L. plantarum isolated from sour-dough cultures produced these compounds and were inhibitory to a number of fungi isolated from baked products (Lavermicocca et al., 2003). The objective of this work was to test L. plantarum and L. paracasei isolated from sourdough bread cultures for antifungal activity against several mycotoxigenic molds and to test the ability of an intact sourdough bread culture to inhibit mold growth.

2.

MATERIALS AND METHODS

The fungi used were as follows: Aspergillus flavus NRRL 1290, Aspergillus ochraceus NRRL 3174, Penicillium verrucosum NRRL 846, Penicillium roqueforti NRRL 848, and Penicillium commune NRRL 1899. Bacterial cultures isolated and identified from the sourdough cultures included, Lactobacillus plantarum 01 and 011, Lactobacillus paracasei 02, 03 and 05 and Lactobacillus paracasei SF1, SF2 and SF21. Isolates 01, 02, 03, 05 and 011 were obtained from an old original household sourdough from West Texas, USA which has been kept active for about 100 years. Isolates SF1, SF2 and SF21 were obtained from a commercial San Francisco type sourdough culture. Sourdough starter cultures and other bacterial isolates were screened for antifungal activity using a dual agar plate assay in which 1.0% of the activated sourdough culture or isolate was added to 15 ml of wheat flour hydrolysate agar (WFH) formulated according to Gobbetti (et al., 1994b), commercial deMan-Rogosa-Sharpe (MRS) agar (Oxoid, Cat. CM0361) and modified MRS (mMRS) agar in Petri dishes. Modified MRS agar was produced by making it without sodium acetate. The sourdough and bacterial isolate agar cultures were overlaid with soft (0.75% agar) yeast extract sucrose agar (YES) or potato dextrose agar (PDA). The centres of the YES or PDA agars were then inoculated at a single point with a mould spore suspension containing 103 spores. The plates were incubated at 30°C for 21 days. Colony diameters of the growing mould cultures were measured in mm and recorded daily. After initial studies the work concentrated on using L. plantarum 01 and L. paracasei SF1 as they appeared to be the most active antifungal

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cultures and were representative of the main isolates. These cultures were then grown and compared on MRS and mMRS agars with PDA overlay.

3.

RESULTS AND DISCUSSION

The influence of bacterial cultures on fungal growth (colony diameter) was plotted as a functional of time (Figures 1-5). Fungal cultures growing on an underlay of uninoculated bacterial media were used as controls. Growth of Aspergillus flavus was delayed in the presence of L. paracasei SF1 and L. plantarum 01 (Figure 1). L. plantarum was more inhibitory than L. paracasei and caused a greater delay in growth. Both bacterial strains appeared to be more inhibitory when grown on MRS agar than mMRS agar, although there was no apparent difference in the growth of the control fungal cultures on the two media. Aspergillus ochraceus was inhibited to a greater degree than was A. flavus (Figure 2). Both L. plantarum and L. paracasei on MRS agar caused complete inhibition of growth of A. ochraceus. On mMRS

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Figure 1. Inhibitory effect of Lactobacillus plantarum O1 and Lactobacillus paracasei SF1 grown in MRS and mMRS agars on growth of Aspergillus flavus NRRL 1290 on a yeast extract sucrose agar overlay.

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Figure 2. Inhibitory effect of Lactobacillus plantarum O1 and Lactobacillus paracasei SF1 grown in MRS and mMRS agars on growth of Aspergillus ochraceus NRRL 3174 on a yeast extract sucrose agar overlay.

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Figure 3. Inhibitory effect of Lactobacillus plantarum O1 and Lactobacillus paracasei SF1 grown in MRS and mMRS agars on growth of Penicillium verrucosum NRRL 846 on a potato dextrose agar overlay.

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agar limited growth of A. ochraceus occurred in the presence of L. plantarum, with growth being delayed until the fifth day of incubation. In the presence of L. paracasei on mMRS growth of A. ochraceus began by the second day of incubation, but growth was limited, though more growth occurred than in the presence of L. plantarum. Growth of Penicillium verrucosum was completely inhibited by L. plantarum on MRS agar and L. paracasei was also more inhibitory on MRS agar than on the mMRS agar (Figure 3). Growth occurred sooner on mMRS, but colonies were smaller than for the control. Penicillium roqueforti grew more rapidly in the presence of the two bacteria than the other mould species (Figure 4). L. plantarium showed a greater inhibitory effect on both MRS and mMRS. P. roquefortii was slightly inhibited by L. paracasei compared with the controls. Growth of P. commune was completely inhibited by L. plantarum on MRS medium and strongly inhibited by L. plantarum on mMRS agar (Figure 5). L. paracasei was less inhibitory than L. plantarum on both media but with no real difference in inhibitory effect between either medium. With all fungal species, differences in growth were seen due to the medium in which the Lactobacillus species were grown. The lactobacilli were more inhibitory when grown in MRS agar than in mMRS. The original MRS medium and the Commercial MRS medium contain 5 g of sodium acetate per litre (0.5%) of medium (de Man et al., 1960). Stiles et al. (2002) reported that L. rhamnosus had greater inhibitory action against 40 different fungal strains when grown in MRS medium than when grown in modified MRS in which the sodium acetate was omitted. They concluded that the acetate was also exerting an inhibitory effect either independently or in addition to inhibitory substances produced by L. rhamnosus. P. roqueforti was least inhibited by either bacterium and did not seem to be inhibited to any greater degree on the MRS over the mMRS by either bacterium, although L. plantarum tended to be more inhibitory than L. paracasei. Penicillium roqueforti is known to have resistance to preservatives such as acetic acid (Gravesen et al., 1994). It appeared possible that MRS may be a better growth medium for lactobacilli from dairy environments, but sourdough cultures may grow better in a medium that has more cereal based ingredients. Therefore, it was decided to add a medium developed by Gobbetti et al. (1994b) called Wheat Flour Hydrolysate Agar (WFH). In this study a complete sourdough mixed culture, not individual bacterial isolates, was added to the base agar layers of MRS, mMRS and WFH in Petri dishes. Base agar layers were then overlaid with YES agar

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Figure 4. Inhibitory effect of Lactobacillus plantarum O1 and Lactobacillus paracasei SF1 grown in MRS and mMRS agars on growth of Penicillium roqueforti NRRL 848 on a potato dextrose agar (PDA) overlay.

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Figure 5. Inhibitory effect of Lactobacillus plantarum O1 and Lactobacillus paracasei SF1 grown in MRS and mMRS agars on growth of Penicillium commune NRRL 1899 on a potato dextrose agar overlay.

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which was inoculated with a single point of A. flavus NRRL 1290 spores in the centre of the plate as previously described. The growth of A. flavus was delayed and reduced on all three media by the intact or complete sourdough culture (Figure 6). Essentially no difference in inhibition was observed in treatments where the sourdough culture was grown in the MRS, mMRS and WFH agars. Thus in this study with the complete sourdough culture there did not appear to be an effect from the medium in which the sourdough culture was grown as was observed with the individual bacterial cultures. Overall this study has shown that L. plantarum and L. paracasei isolated from sourdough bread cultures and an intact sourdough bread culture were inhibitory to several species of mycotoxigenic fungi. The inhibition was manifested both as delay of growth and suppression of the growth rate. The inhibitory effects were influenced by the culture medium or substrate in which the individual sourdough bacteria, but not the complete sourdough culture were grown. The presence of 0.5% sodium acetate in MRS medium resulted in a stronger inhibitory effect by L. plantarum and L. paracasei than mMRS medium from which the sodium acetate was removed. The inhibitory effect was about 20% less when sodium acetate was excluded from the medium. Additional studies are in progress to further evaluate the antifungal

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Figure 6. Inhibitory effect of an intact sourdough bread culture grown in MRS, mMRS and WFH agars on growth of Aspergillus flavus NRRL 1290 on a yeast extract sucrose agar overlay.

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activities of the complete sourdough cultures, and various L. plantarum and L. paracasei isolates.

4.

ACKNOWLEDGEMENTS

This manuscript is published as Paper No. 14941, Journal Series. This research was carried out under Project 16-097, Agricultural Research Division, University of Nebraska-Lincoln and was supported in part by a research grant from the Anderson Research Fund of the NC-213 Multistate Research Project.

5.

REFERENCES

Corsetti, A., Gobbetti M., and Damiani, P., 1998, Antimould activity of sourdough lactic acid bacteria: identification of a mixture of organic acids produced by Lactobacillus sanfrancisco CB1, Appl. Microbiol. Biotechnol. 50:253-256. deMan, J. C., Rogosa, M., and Sharpe, M. E., 1960, A medium for the cultivation of lactobacilli, J. Appl. Bacteriol. 23:130-135. Garver, K. I., and Muriana, P. M., 1993, Detection, identification and characterization of bacteriocin-producing lactic acid bacteria from retail food products, Int. J. Food Microbiol. 19:241-258. Graveson, S., Frisvad J. C., and Samson, R. A., 1994, Microfungi, Munksgaard. Copenhagen, Denmark. Gobbetti, M., 1998, The sourdough microflora: interaction of lactic acid bacteria and yeasts, Trends Food Sci. Technol. 9:267-274. Gobbetti, M., Corsetti, A., and Rossi, I. 1994a, The sourdough microflora: interactions between lactic acid bacteria and yeasts: metabolism of amino acids, World J. Microbiol. Biotechnol. 10:275-279. Gobbetti, M., Corsetti, A. and Rossi, I., 1994b, The sourdough microflora: interactions between lactic acid bacteria and yeasts: metabolism of carbohydrates, Appl. Microbiol. Biotechnol. 41:456-460. Gourama, H., 1997, Inhibition of growth and mycotoxin production of Penicillium by Lactobacillus species, Lebensm. Wiss. Technol. 30:279-283. Gourama, H., and Bullerman, L. B., 1995, Antimycotic and antiaflatoxigenic effect of lactic acid bacteria. A Review, J. Food Prot. 57:1275-1280. Haikara, A., and Laitila, A., 1995, Influence of lactic acid starter cultures on the quality of malt and beer, in: Proceedings of the 26th Congress of the European Brewers Convention, Brussels, IRL Press, Oxford, UK, pp. 249-256. Haikara, A., and Niku-Paavola, M., 1993, Fungicidic substances produced by lactic acid bacteria, FEMS Microbiol. Rev. 12:120. Haikara, A., Uljas, H., and Suurnakki, A., 1993, Lactic starter cultures in malting: a novel solution to gushing problems, in: Proceedings of the 25th Congress of the European Brewers Convention, Oslo, IRL Press, Oxford, UK, pp. 163-172.

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Holzapfel, W. H., Geisen, R., and Schillinger, U., 1995, Biological preservation of foods with reference to protective cultures, bacteriocins and food grade enzymes, Int. J. Food Microbiol. 24:343-362. Karunaratne, A., Wezenberg, E., and Bullerman, L. B., 1990, Inhibition of mold growth and aflatoxin production by Lactobaciillus spp., J. Food Prot. 53:230-236. Klaenhammer, T. R., 1998, Functional activities of Lactobacillus probiotics: genetic mandate, Int. Dairy J. 8:497-505. Lavermicocca, P., Valerio, F., Evidente, A., Lazzaroni, S., Corsetti, A., and Gobbetti, M., 2000, Purification and characterization of novel antifungal compounds from the sourdough Lactobacillus plantarum Strain 21B, Appl. Environ. Microbiol. 66:4084-4090. Lavermicocca, P., Valerio, F., and Visconti, A., 2003, Antifungal activity of phenyllactic acid against molds isolated from bakery products, Appl. Environ. Microbiol. 69:634-640. Niku-Paavola, M.–L., Laitila, A., Mattila-Sandholm, T., and Haikara, A., 1999, New types of antimicrobial compounds produced by Lactobacillus plantarum, J. Appl. Microbiol. 86:29-35. Plockova, M., Stiles, J., and Chumchalova, J., 2000, Evaluation of antifungal activity of lactic acid bacteria by the milk agar plate method, Czech Dairy J. 62:19-19. Plockova, M., Stiles, J., Chumchalova, J., and Halfarova, R., 2001, Control of mould growth by Lactobacillus rhamnosus VT1 and Lactobacilus reuteri CCM 3625 on milk agar plates, Czech J. Food Sci. 19:46-50. Rocken, W., 1996, Applied aspects of sourdough fermentation, Adv. Food Sci. 18: 212-216. Sandine, W. E., 1996, Commercial production of dairy starter cultures, in: Dairy Starter Cultures, T. M. Cogan and J.-P. Accolas, eds., VCH Publishers, New York, pp. 191-206. Spicher, G., 1983, Baked goods, in: Biotechnology, Vol. 5: Food and Feed Production with Microorganisms, G. Reed, ed., Verlag Chemie, Weinheim, Germany. Stiles, J., Plockova, M., Toth, V., and Chumchalova, J., 1999, Inhibition of Fusarium sp. DMF 0101 by Lactobacillus strains grown in MRS and Elliker broths, Adv. Food Sci. 21:117-121. Stiles, J., Penkar, S., Plockova, M., Chumchalova, J., and Bullerman, L. B., 2002, Antifungal activity of sodium acetate, a component of MRS medium, J. Food Prot. 65:1188-1191.

PREVENTION OF OCHRATOXIN A IN CEREALS IN EUROPE Monica Olsen1, Nils Jonsson 2, Naresh Magan3, John Banks 4, Corrado Fanelli5, Aldo Rizzo6, Auli Haikara7, Alan Dobson8, Jens Frisvad 9, Stephen Holmes10, Juhani Olkku11, Sven-Johan Persson12 and Thomas Börjesson13

1.

INTRODUCTION

This paper describes objectives and activities of a major European Community project (OTA PREV) aimed at understanding sources of contamination of ochratoxin A in European cereals and related foodstuffs, and the development of strategies to minimise ochratoxin A in the food supply. The project ran from February 2000 to July 2003.

1

National Food Administration, PO Box 622, SE-751 26 Uppsala, Sweden Swedish Institute of Agricultural and Environmental Engineering, PO Box 7033, SE-750 07 Uppsala, Sweden 3 Cranfield Biotechnology Centre, Cranfield University, Barton Road, Silsoe, Bedfordshire MK45 4DT, UK 4 Central Science Laboratory, Sand Hutton, York YO41 1LZ, UK 5 Laboratorio di Micologia, Univerisità “La Sapienza”, Largo Cristina di Svezia 24, I-00165 Roma, Italy 6 National Veterinary and Food Res. Inst., PO Box 45, FIN-00581, Helsinki, Finland 7 VTT Biotechnology, PO Box 1500, FIN-02044 Espoo, Finland 8 Microbiology Department, University College Cork, Cork, Ireland 9 BioCentrum-DTU, Building 221, DK-2800 Kgs. Lyngby, Denmark 10 ADGEN Ltd, Nellies Gate, Auchincruive, Ayr KA6 5HW, UK 11 Oy Panimolaboratorio-Bryggerilaboratorium AB, P.O. Box 16, FIN-02150 Espoo, Finland 12 Akron maskiner, SE-531 04 Järpås, Sweden 13 Svenska Lantmännen, Östra hamnen, SE-531 87 Lidköping, Sweden. Correspondence to: [email protected] 2

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The Objectives

The over-all objective for the OTA PREV project is the protection of the consumer’s health by describing measures for decreasing the amount of ochratoxin A in cereals produced in Europe. This has been achieved by identifying the key elements in an effective HACCP programme for ochratoxin A for cereals, and by providing tools for preventive and corrective actions. A summary of the tools provided by this project is presented in Table 1. The project included 11 work packages covering the whole food chain from primary production to the final processed food product (Table 2). The objectives and expected achievements were divided into four tasks, all important steps in a HACCP managing programme for ochratoxin A in cereals: 1. Identification of the critical control points (CCP); 2. Establishment of critical limits for the critical control points; 3. Developing rapid monitoring methods, and 4. Establishment of corrective actions in the event of deviation of a critical limit. The outcome will serve as a pool of knowledge for HACCP-based management programmes, which will increase food safety and support the European cereal industry.

1.2.

Why Ochratoxin A?

EC legislation and Codex Alimentarius are currently addressing the problem of ochratoxin A in food commodities and raw materials. Ochratoxin A can be found in cereals, wine, grape juice, dried vine fruits, coffee, spices, cocoa, and animal derived products such as pork products. The current EC legislation includes unprocessed cereals, cereal products and dried vine fruits (Commission regulation 472/2002) and limits for other commodities are being discussed, including baby food. JECFA (the FAO/WHO Joint Expert Committee on Food Additives) evaluated ochratoxin A at its 56th meeting in 2001 (JECFA, 2001). Ochratoxin A is nephrotoxic in all tested animal species and may cause renal carcinogenicity, but the mechanism of action is still being debated. Both genotoxic and non-genotoxic mechanisms have been proposed. JECFA retained the previously established provisional tolerable weekly intake (PTWI) at 100 ng/kg bodyweight (b.w.), corresponding to approximately 14 ng/kg b.w. per day. Estimates of tolerable daily intake for ochratoxin A, based on non-threshold mathematical modelling approaches or a safety factor/threshold approach, have ranged from 1.2 to 14 ng/kg b.w. per day. The Scientific Committee for Food of the European Commission (SCF, 1998) considered that “it would

Buffer storage before drying and during drying (in near-ambient dryers)

CCP

Storage

Intake at cereal processing industry

(up to 100 % prevention possible) Monitoring tools: LFDs and ELISAs.

GSP/ CCP

Recommendations on silo design and maintenance. (WP5) Critical limits for remoistening. (WP5) Food grade antioxidants and natural control measures to prevent OTA formation in wet grain. (WP 6)

(% prevention not possible to estimate, but significant) (up to 100 % prevention possible) (>80 % prevention but not yet economically feasible)

CCP

Rapid monitoring methods for OTA* and OTA producing fungi in grain. (WP8) Critical limit: less than 1000 cfu/g P. verrucosum in wheat. (WP4) Monitoring method for P. verrucosum. (WP1, WP8, and WP9)

LFD (with reader for ochratoxin A) and ELISA

GMP

Reductive measures during milling. (WP10)

(% prevention not possible to estimate, but useful tools in DSS)

Indicating risk of OTA levels above 5 µg/kg Monitoring tools: DYSG, LFD, ELISA, and PCR

(cleaning 2-3%, scouring 3-44%, milling up to 60%) Continued

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Milling industry

Mathematical model which can predict safe storage time (critical limits). (WP4) Rapid monitoring methods for OTA and producing fungi. (WP8) Data on environmental conditions conducive to growth and OTA production. (WP3)

Prevention of Ochratoxin A in Cereals

Table 1. Summary of tools to prevent ochratoxin A in the cereal production chain as provided by the EC project known as OTA PREVa Control Site type Tools provided Comments (possible % reduction of OTA) Harvest GAP Recommendation: Keep machinery and areas in (% prevention not possible to estimate, but contact with the harvested grain, clean. Remove old significant) grain and dust. (WP1)

Intake at malting industry

CCP

Critical limits: